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ALTERATION OF CHLOROPHYLL IN PLANTS UPON AIR POLLUTANT EXPOSURE Robert L. Heath Department of Botany and Plant Sciences University of California Riverside, CA 92521 ABSTRACT The loss of chlorophyll from plants (chlorosis) has been long used to assess injury induced by varied air pollutants, and is believed to signal a decline in photosynthetic capacity and plant productivity. Unfortunately, chlorosis is not specific for air pollutant stress; other stresses and, even, normal development can alter pigment content. In fact, it is not even clear that a loss of chlorophyll consistently mirrors a loss of photosynthetic capacity. This paper will briefly review some concepts of pigment loss, including photosynthetically-linked mechanisms, and the development of pollutant injury. Recently, chlorophyll fluorescence has been touted as a useful monitor of injury and so, some discussion on the methods and instrumentation of chlorophyll measurements, including the use of chlorophyll fluorescence as a monitor, will be presented. CHLOROPHYLL CONTENT Often in stress literature, the terms necrosis and chlorosis are intermixed due to their joint use in monitoring in viva alterations in the leaf. Necrosis often means chlorosis in essence, since chlorophyll molecules in dead cells, when exposed to both light and oxygen in the absence of protestants, are chemically modified and lose visible light absorbance, and so bleach. Chlorosis, however, only means "yellowing caused by loss of or reduced development of chlorophyll... (emphasis added)"~191. In many cases, it is difficult to know to which term the research is referring. For the most part, only visible injury, which includes both chlorosis and necrosis, is reported. Only a portion of the research reported actually measures chlorophyll loss by extracting the pigments and measuring the decline in total extractable pigments. Chlorophyll is porphyrin derivative (a flattened ring of four pyrrole rings) with a fifth ring (the pentanone ring) attached to pyrrole ring III and the r-methylene bridge, with a central Mg ion (Fig. 1~. The chlorophyll molecule is made hydrophobic by the attachment of a phytol chain (tetramethyl hexadecenol) to ring IV (at the 7b carbon) [21. Chlorophyll a and b differ by only one group, a methyl or formyl group on the 3b carbon of ring II (Fig. 1~. Chlorophyll can be easily transformed by the loss of the central Mg ion, caused by acidification, to pheophytin (a distinctly yellow-green pigment), while the loss of phytol, caused by chlorophyllase, produces chlorophyllide with a slightly different spectra . The 6d carbon in the cyclopentone ring can easily isomerize in alcoholic solutions to form allo-chlorophyll, whose spectra is virtually identical to chlorophyll. In the presence of oxygen, various carbons in the pentone ring can be oxidized to form several poorly defined oxidation products. Chlorophylls, of course, have distinctive spectral characteristics. The blue band of the spectra (the Soret Band with an absorbance about 400 nm) overlaps with carotenoid spectra; however, the red band (alpha-band) can be used to separate chlorophyll a from chlorophyll b t2,3,71. Generally, the ratio of chl a/chl b is about 2.5 and so the 347

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348 H Cal 5 ~ ~ ~ N=~/4 a Mg art 8a~N N~5a 8C ~cCH3 ~ ~ 7C 6d ~ 7>,: 1 =0 3 ~ H 1 C. LH H CH3 CH3 H CHIN Figure 1. Structure of Chlorophyll Molecule. Adapted from Aronoff [21. The structure shown is chlorophyll A; the replacement of the methyl group on carbon 3b, with a formyl group (as shown by the insert and arrow) changes the molecule into chlorophyll B. The hydrophobicity of the molecule is provided by the phytol chain shown on the bottom and attached through carbon 7e. separation of individual chlorophyll peaks can be accomplished by a good spectrophotometer. Yet the determination of ratios greater than 3 are generally inaccurate. Also some of the oxidative species possess spectra which are slightly shifted and so can confound the determinations of pure chlorophylls by spectrophotometric methods (see ref. 3,4,12,26,27,30 for some of the peaks and extinction coefficients of chlorophylls and their products in varied solvents). The extraction of chlorophylls by organic solvents can be accomplished to a varied degree. Acetone (80%) is easy and traditional t12,29], ethanol is adequate and often used [4,30], but a series of organic solvents (chloroform + methanol), used for general lipid extraction, perform the best in fully extracting the pigments and their products [4,21,261. Some chemical alterations of chlorophylls can be observed by spectral analysis (however, one should use a relatively sophisticated spectrophotometer); others require chromatography techniques to discover. Examples of the oxidative conversions is shown in Figure 2, in which pigments are extracted through lipid extraction techniques (by the use of chloroform and methanol) and subjected to thin layer chromatography (TLC). After separation, the pigments are rapidly scraped from the plate and extracted from the silica by chloroform and separated by high pressure liquid chromatograph (HPLC). Clearly, other chlorophyll-like compounds have been induced by air exposure through TLC. Some of these extra bands are allomerized chlorophyll [27] and others are oxidized species (see ref [4,21,26] for a more complete discussion of oxidations, spectra and HPLC separations).

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349 In jec1 1 ~ 1 1 0 5 1 1 MA 0.02 Absorption or ~ ' ~ ~ _ _ , _ _ ~ 1 I 10 15 20 25 Tl M E (min) Figure 2. The Modification of Chlorophyll Pigments upon Exposure to Air During Thin Layer Chromography Separation. Pigments from spinach were isolated by standard lipid extractions using chloroform:methanol (1:1, v/v) and the Folch phase separation. The pigment fraction was separated from the polar lipids by a Sep-pak (Silica, from Waters, Inc.) removing ,B-carotene by chloroform wash and the pigments by a chloroform:ethyl acetate (4:6, v/v) wash. The resulting fraction was concentrated by flash evaporation and boiling down under nitrogen and was resuspended in a small volume of chloroform. The concentrated fraction was either run on a C-S HPLC column (5 Am pore size, 4.5 x 150 mm, Beckman; solvent of 80:10: 10, methanol/chlorform/water at a flow rate of 1.5 ml/min) or spotted on a TLC plate (Silica HF plate, 100 m thickness) and developed by a solvent of diethy} ether. The figure shows HPLC runs (injected volume, 20,u1 two regions of the TLC plate (top, chlorophyll A; middle, chlorophyll B) after scrapping, eluting with chloroform, and filtering through a 0.2,um filter. The bottom run shows the pigments fraction that was not run on the TLC plate.

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350 This figure also demonstrates the power of HPLC for quantitifying pigments. In one determination using an isocratic solvent which is complete in 25 minutes, carotenoids and chlorophylls can be quantitated, although the polar carotenoids are not well separated. To separate these and to allow f-carotene to be determined in a single trial, a solvent gradient or several trials with differing solvents of varying hydrophobic characteristics must be used, thus lengthening each analyses. Not shown in this diagram but tabulated in Table 1, tocopherol and its quinone can also be separated on this HPLC system. Table 1. Polarity of Solvent System Used on HPLC and the Relative Retention Times of Several Lipid Components. RETENTION TIME (RELATIVE) Solvent (methanol/chloroform + 10 water) Component 60/25 65/15 70/20 80/10 90/10 ,6-carotene 3.35 Chloro A 1.94 2.54 3.22 5.25 17.0 Chloro B 1.42 1.73 2.10 3.25 9.95 Tocopherol 1.08 2.85 Tocoquinone 0.86 1.92 Lutein 0.47 0.70 1.18 3.63 Neoxanthin 0.26 0.35 0.55 1.50 Vioxanthin 0.12 0.21 0.44 1.24 The retention times of the indicated components were made relative to the retention time of the void volume of the column (see legend in Figure 2~. Solvent for development of HPLC was made as indicated + 10 volumes water, as v/v. Chlorophyll can also be quantitified in vivo to a certain extent, by the use of an integrating sphere [71 (such a system can be purchased through LiCor, model LI- 1800~. When light passes through a leaf, it is highly scattered by the material within the leaf. If the leaf-scattered light is collected after multiple re-scattering, the loss of light by only true absorbance can be determined [71. This system is not perfect and spectra thus derived still possess a large spectral broadening which confounds the actual chlorophyll absorbance. Yet the chlorophyll peak can be observed and quantified. More importantly, this determination is not destructive. However, the ratio of chl a/chl b cannot be determined. Thus, three methods can be used to determine chlorochvll: ( 11 in vivo re-scatterin absorbance determination; or organic solvent spectrophotometric analysis; or (3) HPLC analysis. ~ ~ ~ , ~ extraction followed by (2) Number 1 is the fastest and non-destructive but is the most inaccurate. Both numbers 2 and 3 destroy the plant material. Number 2 is relatively fast but cannot determine all types of pigments accurately. With automatic equipment (which is expensive), number 3 is the most accurate and inclusive. As previously stated, chlorophyll loss is generally related to damage by air pollutants. Len~zian and Unsworth [ 18] believe that sulfur dioxide leads to lysis of chloroplasts and then destruction of the chlorophyll, but only in the "marginal and intercostal areas". Hallgren [91 suggested that the loss of pigments for Pinus contorta (using sulfur dioxide in solution) was caused by the pollutant inducing a solubilization of enzymes from the membrane (in particular, chlorophyllase) which formed chlorophyllids, leading to a loss of chlorophyll from the grana membranes. Ziegler [31 ] suggested sulfur dioxide fumigation of lichen caused the formation of pheophytin (presumably by acidification and monitored by both bleaching and a spectral shift) but stated that chlorophyll loss was too slow to cause the measured decline in photosynthesis. Hallgren

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351 [9] suggests that much of the chlorophyll loss is initiated by pollutant-induced free radicals ted. Certainly, this is plausible in chemical systems where fatty acid hydroperoxides (which break down and form free radicals) can initiate, and tocopherol and carotene (which are antioxidants and free radical chain interruptors) can inhibit chlorophyll bleaching. However, rather than a direct cause of pollutant or its products upon chlorophyll, Davison and Barnes [51 suggested that stress may "pre-dispose the leaf to photooxidation" and "swamp" the photoprotective mechanisms. Many of their experiments involved the combined action of cold and air pollutants in conifers. Frost inhibited photosynthesis and the secondary pollutant stress induced a photooxidation of the structure of the chloroplast. Knudson et al. t15] exposed young pinto bean seedlings to 1 hour of 0.9,1/L v/v) of ozone and followed to the chlorophyll loss by extraction in ethanol for 2, 4, and 6 days post-fumigation. They used visual examination to determine "chlorosis" or '"visible injury" before extraction. They found a close 1:1 relationship between a visually scored plant injury and a loss of extractable chlorophyll. A typical range of visual scoring for a 10% reduction of chlorophyll was about 6 + 6 % while for 50% chlorophyll reduction, the range was 55 ~ 18%. There was no indication in this experiment that chlorophyll a was more or less senstive that chlorophyll b. Thus, the visual scoring monitored general chlorophyll loss and it can be carried out in the field with a minimum of effort. In this experiment, the chlorophyll extraction was likewise easily done by allowing the leaves to sit in ethanol for several days in the dark, but long term ethanolic extraction can have problems with multiple product formation. Lower levels of pollutants may cause a premature senescence rather than a direct chlorophyll loss. Adedipe et al. [1] found in several varieties of tobacco that exposure to 0.31/L ozone (for 2 hours) induced chlorophyll, RNA, and protein loss which followed normal senescence patterns. On the other hand, Leffler and Cherry [17] suggested that the site of the predisposition of injury was the chloroplasts, since reduction of the activity of nitrate reductase (found within the chloroplast) and of the amount of chlorophyll were identical upon ozone fumigation of soybean. On the basis of chemical studies and the results of Sakaki et al. [28], it has been suggested that chlorophyll a is less stable than chlorophyll b and so the increase in the ratio of a/b is a good indication of free-radical-induced pollution effects. Rabe and Kreeb [22] do not believe this, however, based upon their study with ambient air in Stuttgart; they did observe a large increase in the levels of pheophytin within the injuried plants, especially spruce and turnip. Several conclusions can be made: ( 1 ) Chlorophyll loss is found to develop in leaves exposed to oxidants or other air pollutants (such as sulfur dioxide and oxides of nitrogen), yet the loss of chlorophyll takes several days to be observed, and presumably requires light. (2) Chlorophyll loss can be measured with accuracy only when the episode is somewhat severe. Mild exposure may cause chlorophyll changes, but they may be more connected with abnormal states of development (e.g., early senescence or partially closed stomates). (3) While one would expect other products of chlorophyll oxidations to appear, only pheophytin has been observed. Yet a concerted effort to measure products has not yet been undertaken, even in chemical systems. (4) While changes in (chlorophyll a)/(chlorophyll b) ratios are suggested by some to be a monitor of injury, the results are not yet compelling and difficult to observe accurately. (5) Chlorophyll loss is a relatively simple monitor of leaf changes and should be

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352 used for monitoring air pollution injury, but only in relation to other parameters (for example, protein levels or total cell mass) if realistic conclusions wish to be drawn. CHLOROPHYLL FLUORESCENCE Schreiber et al. [25] used chlorophyll fluorescence in viva to monitor ozone-induced stress. Chlorophyll upon absorption of light will naturally fluoresce if a polar solvent is present. The fluoresecence has a life time of about 15 nano-seconds with a yield of emitted light to absorbed light of about 30%. In dry solvents there is no fluorescence; the light energy is dissipated by non-radiative pathways. Light energy absorbed by both the Soret and red bands is reduced to the level of the red band through non-radiative pathway and, when released by fluorescence, is emitted as a band centering about 682 nm (for solvents in which the red band peaks at 663 nary). In plants, the chlorophyll molecules are found within a slightly polar molecular domain and so they fluoresce with a yield of about 2 to 8% efficiency (and a 5- to 1 0-fold faster life-time). Furthermore, there are at least several molecular domains in which the chlorophylls are located, which shifts the emitted fluorescence wavelength peaks to 684 and approximately 720 nm. At room temperature the 720 nm emission is much less (by a factor of 20 or so) than the 684 nm emission. At liquid nitrogen temperature (77 K), where enzymic mechanisms cannot operate, three bands are observed (684, 693, and 730 nary). Researchers have used the 77 K emission to ascertain electron transport properties in leaves, but this is not practical in field experiments. Under normal conditions fluorescence is not constant but varies with post-illumination time [10,161. The initial level of fluorescence is measured (Fo) under a minimum of amount of illumination time. In general, Fo is the smallest level of fluorescence that can be measured regardless of conditions and must be done within milliseconds of the beginning of illumination. The level then begins to rise immediately after illumination up to ~ new level (Fm = cat 1.15 x Fo), within about 20 to 40 milliseconds, depending upon the light intensity. These changes are thought to be due to only primary photochemistry of Photosystem II of the electron transport pathways [16,25,28] and can only be measured with rapidly responding equipment (certainly not pen recorders). A typical slow kinetic trace is shown in Figure 3. The Fo can not be detected under these conditions. However, after a slight, rapidly-occurring hesitation (or sometimes a slight dip) the fluorescence rises again to a new, maximum observable level (called Fp = cat 3 x Fo) in about 1 second. This level is very sensitive to the illumination history of the leaf and to stress. As one of the first observations of pollutant injury [8, 11 ,251, it tends to be reduced. After several seconds of illumination, the peak lowers with the possibility of a shoulder on the decline side occurring at about 5 to 10 seconds. The final steady-state level occurs at about 50 to 100 seconds of illumination ("T" level), and is slightly above Fo. Herbicides affect this fluorescence kinetic curve dramatically and help understand what the kinetics mean. Methyl viologen (paraquat, MV) can be infiltrated into leaf discs and by accepting electron at the reducing side of Photosystem I (before NADP reduction), inhibits carbon dioxide fixation. By taking electrons rapidly from the electron transport pathway, MY keeps the Photosystem largely oxidized and, as is seen in the figure, keeps the kinetics of fluorescence at a low level (near Fo). Dichlorophenyl dimethyl urea (DCMU) blocks the exit of electrons out of Photosystem II and keeps Photosystem I largely oxidized and Photosystem II largely reduced. DCMU also inhibits carbon dioxide fixation and keeps the fluorescence level very high with little kinetic change. The DCMU-induced level is nearly the highest fluorescence level that can be seen under any conditions. Thus, MV and DCMU can induce the minimum level (Fo) and the maximum level (Fmax) of fluorescence, respectively. The range between these two levels is often denoted as the variable fluorescence (Fv).

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353 A water stress can be applied to leaf discs by floating them on a concentration solution of Polyethylene Glycol 6000 (PEG), which osmotically removes water from the tissues. The fluorescence pattern is modified by this treatment All, as shown in the figure. The peak is lowered, which accentuates the shoulder at about ~ seconds. CONTROL if me, ~ ~ 1 1 +MV + DCMV . _ L o 40 80 o 40 80 o 40 T I ME I N SECONDS + PEG - 1 1 1 1 1 80 o 40 80 Figure 3. In Vivo Fluorescence Kinetics of Spinach Leaf Disc. Discs from spinach leaves were cut and floated on water- solutions for 1 hour in the dark, as pre-conditioning. The fluorescence data (excited by 400-450 nm light and observed at 680-690 nm) were obtained from a Hansatech Oxygen-/fluorometer [28 and see text]. The vertical scale is relative but constant. The water solutions were (Atwater alone (control), (b) methyl viologen (100pm,mv), (c) Dichlorophenyl dimethyl urea (30 m,DC), and (b) polyethylene glycol 6000 (12%, PEG). Furthermore, ~ new peak can be seen at 60 seconds (the M peak), which is thought to be a modification of the light energy flow within the chloroplast (see 13,16,28~. Ozone injury resembles this pattern (see Schreiber et al., 25~. A lowering of Fo is only observed when gross injury to the system has occurred and most probably great chlorophyll loss is likewise occurring [11,25~. The immense body of literature on chlorophyll fluorescence and its interpretation (for some reviews, see [8, 1 0, 11 , 1 3, 1 4, 1 6,20,24,281), cannot be summarized here. It is possible to monitor fluorescence relatively easily now, even in the field. The interpretation of fluorescence kinetics alterations due to stress, however, is much more difficult. ~ The above description of the kinetics represents only the direct illumination and emission type of measurements [ 103. Another type of instrument involves the direct measurement of yield and is independent of illumination [20~. A constant, weak measuring beam of light (which is modulated for ease of detection) illuminates the leaf and the detector is "locked" onto the modulated, emitted fluorescences, which is in phase

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354 with the illumination. Other direct, non-modulated light instruments, but will excite the photochemistry of the leaf one of a base level (near Fo), and its rise to extra-illumination. When the extra light is turned off, level again (see Figure 4~. Again the Fp is sensitive to for a preliminary experiment with Jeffery Pine (from G. Randel, UCLA). \ will not be detected by the Thus, the measurement is a higher level (Fp) due to the yield declines to the base stress, as shown in the figure Goldstein, Argentine, and P. \ ~ min m a Figure 4. The Fluorescence Yield of Pine Needles. Data provided by G. Goldstein and P. Randel. Needles from Jeffrey Pine trees exposed to ambient conditions in the Los Angeles basis were measured on the yield fluorometer [13 and see text]. The measuring light is turned on at m and the actinic white light (intensity of 2000pm-teins/cm2 see) is turned on at a. The bottom panel shows four year old needles from non-injuried (solid line) and severely injuried (dotted line) trees. Thus, fluorescence can monitor the reactions occurring within the chloroplast. The changes of the pattern can be easily detected currently but the understanding of what is occurring is less easy to come by, regardless of what some believe. Certainly, under mild stress, fluorescence may turn out to be a reasonable monitor. However, under very mild stress (0.24 zone for 4 hours under low humidity conditions), reproducible changes in Fp, in spinach and bean, have not been observed, even when the yield of dry matter was reduced 10-20% (unpublished data of D. Olszyk and R.L. Heath). It is clear that more basic work needs to be done before chlorophyll fluorescence is a simple monitor of photosynthetic alterations by pollutants. CONCLUSION While visible injury still dominates the literature as a marker of pollutant injury, the technology is available to measure chlorophyll loss in viva and to monitor some alterations of the primary events of the chloroplasts through chlorophyll fluorescence kinetics. However, it is too soon to be able to interpret these changes biochemically and these alterations may not be specific to only air pollutant injury. Indeed, it is doubtful that a specific monitor of air pollutant injury will ever be discovered. More probably, several markers must be correlated before it is certain that only air pollutants are the cause of a plant's stressful state.

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355 REFERENCES 1. Adedipe, N.O., R.A. Fletcher, and D.P. Ormrod. 1973. Ozone Lesions in Relation to Senescence of Attached and Detached Leaves of Tobacco. Atmosph. Environ. 7: 357-361. 2. Aronoff, S. 1966. The Chlorophylls--An Introductory Survey. In: The Chlorophylls. (Eds., L.P. Vernon and G.R. Seely) Academic Press, New York, Pp. 3-21. 3. Bruinsma, J. 1961. A Comment on the Spectrophotometric Determination of Chlorophyll. Biochim. Biophys. Acta 52: 576-578. 4. Braumann, T., and L. H. Grimme. 1981. Reversed-Phase High Performance Liquid Chromatography of Chlorophylls and Carotenoids. Biochim. Biophys. Acta 637: 8-17. 5. Davison, A.W., and J.D. Barnes. 1986. Effects of Winter Stress on Pollutant Responses. In: How are the Effects of Air Pollutants on Agricultural Crops Influenced by the Interaction with Other Limiting Factors? Proc. of Workshop, Commission of European Communities and Natl. Agency of Environ. Protect., Denmark, pp.l6-32. 6. Saran, M., C. Michel, and W. Bors. 1988. Reactivities of Free Radical In: Air Pollution and Plant Metabolism (Eds., S. Schulte-Hostede, N. Darrall, L. Blank, A.R. Wellburn), Elsevier Publishers, London Pp. 76-93. 7. French, C.S. 1960. The Chlorophylls in Vivo and in Vitro. In: Encycl. of Plant Physiology. (Ed. W. Ruhland) Vol. V/1, Springer-Verlage, Berlin, pp. 252-297. 8. Govindjee, W.J.S. Downton, D.C. Fork, and P.A. Armond. 1981. Chlorophyll A Fluorescence Transients as an Indicator of Water Potential of Leaves. Plant Sciences Lett. 20: 191-194. 9. Hillgren, J.E. 1979. Physiological and Biochemical Effects of Sulfur Dioxide on Plants. In: Sulfur in the Environment Part II. Ecological Impacts. (Ed. J. O. Nriagn). John Wiley and Sons, New York. Pp. 164-209. 10. Heath, R.L. 1973. The Energy State and Structure of the Isolated Chloroplast: The Oxidative Reactions Involving the Water Splitting Step of Photosynthesis. Intl. Rev. Cytol. 34: 49-101. 11. Heath, R.L., P.E. Frederick, and P.E. Chimiklis. 1982. Ozone Inhibition of Photosynthesis in Chlorella Sorokiniana. Plant Physiol. 69: 229-233. 12. Inskeep, W.P., and P.R. Bloom. 1985. Extinction Coefficients of Chlorophyll a and b in N,N-Dimethylformamide and 80% Acetone. Plant Physiol. 77:483-485. 13. Jenkins, G.I., N.R. Baker, M.Bradbury, and H. W. Woolhouse. 1981. Photosynthetic Electron Transport During Senescence of the Primary Leaves of Phaseolus vulgaris L. III. Kinetics of Chlorophyll Fluorescence Emission from Intact Leaves. J. Expt. Bot. 32: 999- 1008. 14. Krause, G.H., and E. Weis. 1984. ChlorophyI1 Fluorescence as a Tool in Plant Physiology. II. Interpretation of Fluorescence Signals. Photosynth. Res. 5: 139-157. 15. Knudson, L.L., T.W. Tibbitts, and G.E. Edwards. 1977. Measurement of Ozone Injury by Determination of Leaf Chlorophyll Concentration. Plant Physiol. 60: 606-608. 16. Lavorel, J., and A.-L. Etienne. 1977. In Vivo Chlorophyll Fluorescence. In: Primary Processes of Photosynthesis (Ed., J. Barber) Elsevier/No. Holland, London. Pp.206-268.

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356 17. Leffler, H.R., and J.H. Cherry. 1974. Destruction of Enzymatic Activities of Corn and Soybean Leaves Exposed to Ozone. Can. I. Botany 52: 1233-1238. 1 S. Len~zIan, K.J., and M.H. Unsworth. 1983. Ecophysiological Effects of Atmospheric Pollutants. In: Encycl. of Plant Physiol. New Series. (Eds., D.L. Lange, P.S. Nobel, C.B. Osmond, H. Ziegler) Vol. IV. Springer-Verlag, Berlin. Pp.466-502. 19. Little, R.J., and C.E. Jones. 1980. A Dictionary of Botany. Van Nostrand/Reinhold Co., New York. Pp. 80. 20. Ogren, E., and N.R. Baker. 1985. Evaluation of a Technique for the Measurement of Chlorophyll Fluorescence from Leaves Exposed to Continuous White Light. Plant, Cell, Environ. 8: 539-547. 21. Prenzel, U., and H.K. Lichtenthaler. 1979. Separation of Prenyllipids by High Peformanance Liquid Chromatography. In: Adv. in Biochem. and Physiol. of Plant Lipids. (Eds., L.-A. Appelqvist and C. Lijenberg), Elsevier/No. Holland, New York. Pp. 391-325. 22. Rabe, R., and K.H. Kreeb. 1980. Bioindication of Air Pollution by Chlorophyll Destruction in Plant Leaves. Oikos 34:163-167. 23. Sakaki, T., N. Kondon, and K. Sugahara. 1983. Breakdown of Photosynthetic Pigments and Lipids in Spinach Leaves with Ozone Fumigation: Role of Active Oxygens. Physiol. Plant. 59: 28-34. 24. Schreiber, U. 1983. Chlorophyll Fluorescence as a Tool in Plant Physiology. I. The Measuring System. Photosynth. Res. 4: 361-373. 25. Schreiber, U., W.Vidaver, V.C. Runeckles, and P. Rosen. 1978. Chlorophyll Fluorescence Assay for Ozone Injury in Intact Plants. Plant Physiol. 61:80-84. 26. Shioi, Y., R. Fukae, and T. Sasa. 1983. Chlorophyll Anaysis by High-Performance Liquid Chromatography. Biochim. Biophys. Acta 722:72-79. 27. Vernon, L.P. 1960. Spectrophotometric Determination of Chlorophylls and Pheophytins in Plant Extracts. Analyt. Chem. 32:1144-1150. 28. Walker, D.A. 1987. The Use of the Oxygen Electrode and Fluorescence Probes in Simple Measurements of Photosynthesis. Oxygraphics Limited Publishing, Chichester, and Hanstech Limited, Norfolk, England. 144 Pp. 29. Wellburn, A.R., and H. Lichtenthaler. 1984. Formulae and Program to Determine Total Carotenoids and Chlorophyll A and B of Leaf Extract in Different Solvents. In: Advances in Photosynthesis Research. (Ed. C. Sybesma) Vol. II, Martinus Nijhoff/ Junk Publishers, the Hague. Pp. 1.9-1.12. 30. Wintermans, J.F.G.M., and A. DeMots. 1965. Spectrophotometric Characteristics of Chlorophyll a and b and their Pheophytins in Ethanol. Biochim. Biophys. Acta 109: 448-453. 31. Ziegler, I. 1975. The Effect of Sulfur Dioxide Pollution on Plant Metabolism. Residue Reviews 56: 79-105.