5 to a higher energy
1 conversion to heat
length. In plants,
te response of plants
lucrescence and the
d near infrared (740
) 650 nm. Another
diation. The origin
iated with both non-
)f the plant. These
ir determination by
lliams 1987; Guyot,
itation wavelengths
ns drive a complex
transfer an electron
velength photons is
na at 690 and 735
hotosystem 2, while
rbs at 680 nm, so
asured fluorescence
ind Rinderle, 1988).
the reflectance and
Olioso et al. (1989)
)ring plant canopies
cence with maxima
1984; Lichtenthaler
87) used ultraviolet
, 685, and 740 nm.
luorescence spectra,
minated at 337 nm.
lg plant types. The
minor maximum or
ots. The difference
logy and leaf vein
;nt maximum at 525
rapid transfer of the
pelle and Williams,
In a review, Guyot (1993) concluded that blue-green fluorescence is largely independent of photosynthetic
activity, but is affected by some types of stress. However, Chappelle, et al., (1991) showed a strong relationship
between the rate of photosynthesis and the ratio of certain wavelengths in the blue fluorescence. Furthermore, they
speculated that the measurement of the changes in blue fluorescence may prove to be useful as a means of remote
estimating of the rate of photosynthesis. Mineral deficiencies (e.g., N and K) and chemical agents capable of
blocking electron transport (e.g., DCMU) reduced photosynthetic efficiency and caused increases in chlorophyll (red)
fluorescence with little effect on blue fluorescence (Chappelle et al., 1984; McMurtrey et al., 1994). Water stress,
on the other hand, impaired photosynthesis and caused increases in both red and blue fluorescence (Chappelle et al.,
1984). A decrease in chlorophyll concentration in leaves was accompanied by decreases in chlorophyll fluorescence
at 690 and 740 nm and increases at 440 and 525 nm.
The origin of the blue-green fluorescence emission of plants is not clear. Numerous compounds found in
plants have a blue-green fluorescence. Several researchers have suggested that the blue-green fluorescence of plants
might be attributed to bound nicotinamide adenine dinucleotide phosphate (NADPH), lignin, esterified ferulic acids,
phenylpropanes, and other phenolics (e.g., chlorogenic acid and caffeic acid) (Chappelle et al., 1991; Goulas et al.,
1991; Lichtenthaler et al., 1991). Other contributors to the blue-green fluorescence are beta-carotene and vitamin
K, (Chappelle et al., 1991). Stober and Lichtenthaler (1993) demonstrated that the cell walls of the epidermal layer
as well as the cells of the xylem and phloem exhibited a strong blue-green fluorescence. Mesophyll cells in green
leaves showed only red chlorophyll fluorescence indicating reabsorption of the emitted blue-green fluorescence by
the broad absorption bands of chlorophylls and carotenoids (Stober and Lichtenthaler, 1993). Thus, it is probable
that the blue-green fluorescence of a leaf is the sum of the fluorescence of different chemical compounds, some of
which are located in the vacuoles and the cell walls (Lang et al 1991; Lichtenthaler et al., 1991; Stober and
Lichtenthaler, 1993) while others are associated with the photosynthetic mec hanis m in the chloroplasts (Chappelle
et a., 1991; 1993).
2 - MATERIALS AND METHODS
At the ARS research station near Akron, Colorado, field trials were conducted with wheat (Triticum aestivum L.
'Oslo). Shortly after harvest, samples of standing wheat residue were collected, air dried, and cut into 5-10 mm long
pieces. Glass, 155 ml serum vials, containing 50 g silica sand and 1.0 g of wheat residue, were prepared and
weighed. Into each vial, we added 12 ml of buffer (Reinertsen et al., 1984) and 1 ml of supernatant from freshly
collected soil extract. The extract was prepared by suspending 50 g of soil in 50 ml of residue buffer, mixing, and
centrifuging at 500 xgfor5 minutes. All bottles were plugged with foam stoppers and incubated in the dark at
30°C. Deionized water was added to the bottles weekly to adjust for evaporative losses. After 0, 0.5, 1, 2, 4, and
8 weeks of incubation, the botdes were removed from the incubator, dried at 60 C, and weighed. Changes in dry
weight with time were used as a measure of residue decomposition.
The dried wheat residues were ground to pass a 1 mm screen, placed in a quartz cuvet, and excited with
340 nm radiation in a SPEX Fluorolog-2 spectrofluorometer. The emission spectra of each sample was measured
at 5 nm interval over the 360-600 nm wavelength range. A subsample of residue from each date was extracted with
hot methanol and then acetone using procedures described by Chappelle et al. (1991). After each extraction, the
wheat residue was dried and its fluorescence spectra was measured again. Additional samples of residue are being
analyzed for changes in fiber composition, but will not be discussed here. Fluorescence spectra of several
representative soils, identified from the survey of U.S. soils by Daughtry et al. (1993), were also measured.
m
440/685
30
5.40
41
1.96
31
0.53
25
3.83
78
0.09
3 - RESULTS AND DISCUSSION
During the 8 weeks of incubation the wheat residue lost slightly more than 40% of its original weight (Figure 1) and
changed from a golden tan color to a dark brown color. In a survey of the literature cm plant residue decomposition,
Jenkinson (1971) reported that the proportion of crop residue decomposed under different climatic conditions was
remarkably similar. Approximately one-third of the residue remained after 1 year. Under field conditions,
temperature and moisture limit decomposition. Thus decomposition that occurred during the 8 -week incubation under
t^arly optimal conditions used in this study, probably represented 6 to 12 months of decomposition under field
conditions.
The fluorescence emission spectra of the wheat residue at each time are shown in Figure 2. The maximum
for each spectra was at 440 + 5 nm. After 8 weeks, the fluorescence intensity of the residue was less than 30% of
its initial value, but was still significantly greater than the emission of the soils. MuMurtrey et al. (1993) reported
859
