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where long-term observations have shown
that neither sex disperses (16). However,
our data further show that adult males
never father offspring within their home
pods (whether this is also true of killer
whales must await parentage testing). By
implication, pilot whales must mate when
two or more pods meet or when adult males
pay short visits to other pods. Both possi-
bilities are supported by field observations.
Large aggregations of whales have been
reported, sometimes numbering well over
1000 individuals, and probably result from
the temporary merging of several pods (4).
On the other hand, there are also rare,
seasonal sightings of small all-male groups
(4). In either case, the transitory nature of
these events is emphasized by the general
failure of paternity testing to reveal fathers.
This behavior pattem is unusual for mam-
mals. Normally, adult males living in social
groups are expected to maximize their repro-
ductive success by competing for access to
females. This behavior may lead to harem
polygyny, with one or a few dominant breed-
ing males who either force subordinate males
to disperse or prevent them from breeding (2).
It appears that pilot whales neither show
strong reproductive dominance nor disperse
from their natal groups.
The ecology of the pilot whale provides
a possible explanation for this behavior. If
opportunities to mate with females in other
pods are not limited, the optimal male
strategy need not involve caring for his
direct offspring. He might do better by
helping the large number of known rela-
tives in his natal pod (17). However, it is
unclear what benefits a male could provide.
Both defense and assistance in a communal
feeding strategy are possibilities, but they
lack observational support. Whatever the
selective forces involved, it seems clear that
the high degree of relatedness between pod
members can explain to a large extent the
extraordinary cohesion of pilot whale pods.
REFERENCES AND NOTES
1. R. L. Brownell, Jr., and K. Ralls, Rep. Int. Whaling
Comm. 8 (spec. issue), 383 (1990).
2. T. H. Clutton-Brock, Proc. R. Soc. London Ser. B
336, 339 (1989); S. T. Emlen and L. W. Oring,
Science 197, 215 (1977).
3. T. H. Clutton-Brock, Nature 337, 70 (1989); P. J.
Greenwood, Anim. Behav. 28, 1140 (1980).
4. D. E. Sergeant, Bull. Fish. Res. Board Can. 132, 1
5. J. P. Joensen, Ethnol. Scand., 5 (1976).
6. W. Amos, J. A. Barrett, G. A. Dover, Heredity 67,
7. D. Tautz, Nucleic Acids Res. 17, 6463 (1989); C.
Schlbtterer, W. Amos, D. Tautz, Nature 354, 63
8. In four cohorts (n = 6, 3, 3, and 10), all paternal
alleles were different. Pairs of alleles occurred in
two cohorts of n = 6 (one pair each) and one
cohort of n = 12 (two pairs). Of these four pairs,
two were ambiguous because a mother-fetus pair
shared both alleles. These -data support only one
possible instance of a male fathering four off-
spring. The estimated mean, assuming both am-
biguities favored shared paternity, is 1.2 offspring
per male per cohort.
9. W. Amos, Symp. Zool. Soc. London, in press.
10. The empirical mean value of Vwas 0.83.
11. Genotype frequency indices were calculated for an
individual of genotype ab as the square root of
pfapft, where pfa is the pod-specific frequency of
12. In 1000 runs, the mean rvalue (+ SD) was 0.0009
13. We reassigned alleles randomly to "mothers"
(females of sufficient age) with up to nine off-
spring, using segregation ratios determined by
sampling a binomial distribution. In the most ex-
treme case (two-thirds of the pod in nine-offspring
families) slight positive correlations were generat-
ed (mean + SD = 0.082 + 0.11).
14. A female reproduces first at age 5 and thereafter
every 3 years with probability P. Founding and
paternal alleles are selected, with full replacement,
from a alleles each at equal frequency. Simulations
were stopped at target pod size t or when the
oldest female reached age 45. We simulated relat-
ed founders by allowing the founder to reach age
60 and then deleting all individuals >45 years old.
Mean r values varied with P, t, and a (default
values 0.7, 100, and 100, respectively), but all lay
in a range consistent with the empirical value.
15. Following Edwards's method of Support [A. W. F.
Edwards, Likelihood (Cambridge Univ. Press,
Cambridge, 1972)], Support = 13.3 (Leynar) and
9.7 (Midvagur), approximately 6 x 105 and 1.6 x
104 more likely.
16. M.. .Bigg, P. F. Olesiuk, G. M. Ellis, J. K. B. Ford,
K. C. Balcomb vll, Rep. wIt.
(spec. issue), 383 (1990).
17. J. F. Wittenberger, in Handbook of Behavioral
Neurobiology, P. Marler and J. Vandenbergh,
Eds. (Plenum, New York, 1979), vol. 3, pp. 271-
340; E. L. Charnov, Anrim. Behav 29, 631 (1981).
18. We thank G. Desportes, R. Mouritsen, D. Bloch,
and the Faeroese government for sample collec-
tion; D. Bloch and C. Lockyer for analyzing teeth
sections; J. Barrett for statistical advice; and W.
Arnold, F. Trillmich, P. Clapham, J. Pemberton, T.
Clutton-Brock, and an anonymous reviewer for
helpful comments on the manuscript. B.A. was
supported by the National Environment Research
Council and the Royal Society.
Whaling Comm. 12
15 September 1992; accepted 1 February 1993
A Laccase Associated with Lignification
in Loblolly Pine Xylem
Wuli Bao,* David M. O'Malley, Ross Whetten,
Ronald R. Sederoff
Peroxidase has been thought to be the only enzyme that oxidizes monolignol precursors
to initiate lignin formation in plants. A laccase was purified from cell walls of differentiating
xylem of loblolly pine and shown to coincide in time and place with lignin formation and to
oxidize monolignols to dehydrogenation products in vitro. These results suggest that
laccase participates in lignin biosynthesis and therefore could be an important target for
genetic engineering to modify wood properties or to improve the digestibility of forage
In 1933, Erdtman proposed that the final
step in lignin biosynthesis was enzymatic
oxidation of p-hydroxyphenylpropanoid
compounds followed by a free radical cou-
pling reaction (1). Peroxidase (E.C.
188.8.131.52) and laccase (E.C. 184.108.40.206) were
postulated to carry out this oxidation be-
cause both enzymes produce dehydrogena-
tion polymers (DHP), a lignin-like materi-
al, from monolignol precursors (2).
Laccase, in contrast to peroxidase, has
rarely been studied in plants. A role for
laccase in lignification was suggested by
early studies with a fungal enzyme (2) but
was later discounted because a purified
plant laccase from the Japanese lacquer
tree (Rhus vernicifera Stokes) was shown
not to oxidize monolignols (3). Many
researchers have associated peroxidases
with lignification (3-7). In studies of
green ash (Fraxinus pennsylvanica Marsh.)
sapling stems, Harkin and Obst showed by
histochemical staining with syringaldazine
Department of Forestry, P.O. Box 8008, North Carolina
State University, Raleigh, NC 27695.
*To whom correspondence should be addressed.
or furoguaiacin in the presence of H202
that the xylem tissue adjacent to the
cambium contained large amounts of per-
oxidase activity (4). These researchers did
not detect laccase activity when syrin-
galdazine was used as the substrate without
H202, and on this basis they concluded
that peroxidase was the exclusive phenol
oxidase responsible for the dehydrogena-
tive polymerization of lignin precursors.
Further evidence of peroxidase involve-
ment in lignification was provided by the
demonstration of peroxidase activity in
differentiating poplar (Populus x eurameri-
cana) xylem (7) and in lignifying cell walls
of differentiating tobacco (Nicotiana
tabacum L.) xylem (6), although some
phenol oxidase activity was detected in
the absence of added H202 in tobacco
The recent characterization of a laccase
purified from the cell culture medium of
sycamore maple (Acer pseudoplatanus L.)
has prompted a reevaluation of the role of
this enzyme in lignification (8-10). We
undertook a study of oxidative enzymes in
differentiating xylem of loblolly pine (Pi-
SCIENCE * VOL. 260 * 30 APRIL 1993
>:.... .R> W
: .:> .R . ... ........ .: R E PO R T S
nus taeda L.) to establish whether laccase
participates in lignin polymerization and
thereby test the hypothesis of exclusive
Differentiating xylem in conifers is an
excellent system for studies of lignin bio-
synthesis because of the high degree of
specialization in wood formation. Crude
cell wall preparations of loblolly pine xy-
lem can oxidize phenolic compounds in
the absence of added H202, which indi-
cates that polyphenol oxidase activity is
present. There are two types of polyphenol
oxidases in plants, laccases and catechol
oxidases, and they can be distinguished by
substrate and inhibitor specificity, copper
content, and molecular weight (11). We
extracted the pine xylem cell walls with 1
M CaCl2 (12), which releases most of the
polyphenol oxidase activity as well as sub-
stantial amounts of peroxidase activity,
Fig. 1. Silver-stained
showing the purified pine
xylem laccase before
(lane 2) and after (lane 3)
deglycosylation (15). The 30
top band of undenatured
laccase (lane 4) coin-
cides with enzyme activ-
ity, as deduced by staining with 0.68 mM DAF in
20 mM Bistris buffer, pH 5.9. The sizes of molec-
ular markers (lane 1) are given in kilodaltons.
Table 1. Substrate specificity of pine xylem
laccase. Reaction rates were monitored with an
oxygen electrode (YSI model 5300, Yellow
Springs Instrument Co., Yellow Springs, Ohio,
1.7-ml chamber). Reaction mixtures contained
0.3 ,g of laccase and 5 mM substrate in 20 mM
Bistris buffer (pH 5.9). We determined the spon-
taneous oxidation rate by monitoring oxygen
depletion in substrate solutions without en-
zyme. The oxygen content in air-saturated buff-
er at 250C was assumed to be 240 ,M (8). ND,
*The solubility of syringaldazine was too low for detec-
tion of oxygen consumption, so laccase activity was
monitored by the formation of a colored product.
and purified to homogeneity a glycopro-
tein (13) with characteristics of a laccase
(14). Both SDS-polyacrylamide gel elec-
trophoresis (PAGE) (Fig. 1) and C4 re-
versed-phase high-performance liquid
chromatography (HPLC) confirmed the
purity of the protein.
The apparent molecular weight of the
glycosylated pine xylem laccase on SDS-
PAGE was -90,000 and that of the degly-
cosylated protein (15) -70,000 (Fig. 1).
Because the laccase activity was only par-
tially denatured by SDS, the active en-
zyme could be detected on SDS gels. The
isoelectric point of the laccase was -9.0,
and the optimum reaction pH was 5.9.
Concentrated solutions of the laccase were
blue with an absorbance peak at 610 nm,
indicative of type I copper (11). Prelimi-
nary results from electron spin resonance
(ESR) spectra also suggested the existence
of a type I copper site in the laccase (16).
We determined the reaction rates of the
pine xylem laccase with several phenolic
substrates (Table 1) by measuring the rates
of oxygen consumption. The fastest reac-
tion rates were found for phenylhydrazine
(an inhibitor of catechol oxidase) and py-
rogallol. The laccase oxidized syrin-
galdazine and catechol, which are com-
monly used substrates for peroxidase and
catechol oxidase assays, but did not oxidize
cresol, guaiacol, tyrosine, or ferulic acid.
Moderate reaction rates were observed for
the monolignols coniferyl alcohol and
sinapyl alcohol, and a slow reaction rate
was observed with p-coumaryl alcohol.
EDTA and KCN, known inhibitors of lac-
case, reduced the rate of oxygen consump-
tion. Diethyldithiocarbamate (1 mM) had
no effect on oxygen consumption but re-
duced color formation in the spectrophoto-
metric assay (14). The Michaelis constant
(Kin) values (17) for coniferyl alcohol and
sinapyl alcohol were 12 ? 1.3 mM and 25.4
? 5.5 mM, respectively. The Km value
could not be estimated for coumaryl alcohol
because the amount of activity was small
and coumaryl alcohol has low solubility.
When 4-methylcatechol was used as sub-
strate, the K. value for oxygen was 37 ?
Table 2. Composition of DHP made by pine xylem laccase. DHP was produced at 250C in 100 [lI
of 20 mM Bistris buffer (pH 5.9) that contained 0.005% laccase. Monolignol substrate in methanol
was added in 1 0-,ug amounts at 1 0-min intervals for 2 hours. DHP was collected by centrifugation,
dried under vacuum, then resuspended in dimethylformamide (DMF). DHP composition was
analyzed by high-performance gel permeation chromatography (KD-802 column, Millipore, Bed-
ford, Massachusetts) at 500C, with DMF containing 50 mM LiCI and 50 mM NaOH as the mobile
phase. Dichlorophenolindophenol, diaminobenzidine, and two degraded lignin samples (620
daltons and 2700 daltons) were used as standards.
Composition of DHP product (%)
Dimer Trimer Tetramer Pentamer
Flg. 2. Histochemistry of lac-
case and peroxidase in 30-,um
cross sections of fresh pine
stem xylem. The sections are
oriented with the cambial side
to the right and the mature xy-
lem side to the left. (A) Sections
were treated at 250C with cata-
lase (1 mg/ml; Sigma) in 20 mM
tris HCI (pH 8) for 10 min then n
stained with 0.68 mM DAF in 20
mM Bistris (pH 5.9) for 30 min.
Laccase activity was detected
only in lignifying xylem near the
cambium. (B) Section treated
with 1% phloroglucinol (in a
25% HCI:75% ethanol solution),
showing the cell walls contain-
ing lignin (red stain). (C) Higher
magnification view of cell walls in the lignifying zone after staining for laccase. (D) Higher
magnification view of cell walls stained for peroxidase, showing that cell walls both in the lignifying
zone and in the mature xylem have peroxidase activity. (E) Higher magnification view of
phloroglucinol staining, showing the weak staining of nonlignifying cell walls near the cambium. (F)
Boiled control section showing no staining for laccase or peroxidase activity. Size bars equal 160
pm in (A) and (B) and 80 pm in (C) to (F).
SCIENCE * VOL. 260 * 30 APRIL 1993
The pine xylem laccase oxidized mono- Download full-text
lignols in vitro to form DHP (18). The
reaction products were primarily trimers
or tetramers of the monolignol (Table
2), although -8% of the DHP from co-
niferyl alcohol was hexamers or larger
All of the detectable pine xylem laccase
activity was bound to cell walls (Table 3)
and could be attributed to a single protein.
No other polyphenol oxidase activities were
detected in the xylem soluble extract.
Strong laccase activity was detected in dif-
ferentiating xylem and in embryogenic cal-
lus, but no activity was detected in embry-
os, megagametophytes, or strobili. When
crude CaCl2-extracted protein samples
were subjected to electrophoresis on SDS
gels and stained for laccase activity, they
produced one broad band that comigrated
with the purified enzyme, which suggests
that there is only one laccase isozyme in
In sledge microtome sections cut from
fresh blocks of pine stem, cells with lac-
case activity were detected by staining in
0.68 mM diaminofluorene (DAF) in 20
mM Bistris bufer (pH 5.9). Laccase activ-
ity was distinguished from peroxidase by
its requirement for oxygen instead of ex-
ogenously added H202. Some sections
were treated with catalase [1 mg/ml in 20
mM tris HCl (pH 8)1 to remove endoge-
nous H202, but no differences in staining
were observed when sections were not
treated with catalase. Only the walls of
xylem cells in the zone of lignification had
laccase activity (Fig. 2A). Phloroglucinol
staining (Fig. 2, B and E) and ultraviolet
Table 3. Laccase and peroxidase activities in
different pine tissues and organs. Crude protein
was extracted overnight at 400 from 5 g of
frozen, powdered plant material in 1 M CaCI2
and 20 mM tris HCI (pH 8) and dialyzed. Xylem
cell wall salt extract was obtained from the
CaCI2 extraction step in the laccase purifica-
tion. Xylem soluble extract was the 10,000g
supernatant from the first extraction of pow-
dered frozen xylem. Laccase activity was de-
termined by spectrophotometric assay (14).
ND, not detected.
Phloem and cambium
Xylem soluble extract
Xylem cell wall salt
fluorescent microscopy indicated the loca-
tion of cells that were initiating lignifica-
tion. Intense laccase activity staining was
observed in the active lignifying zone but
not in mature xylem (Fig. 2, A and C).
Syringaldazine staining of sections in the
absence of added H202 showed a pattern
similar to that in the DAF-stained sec-
tions. Sections boiled in water showed no
laccase activity (Fig. 2F). The addition of
50 mM EDTA to the stain solution inhib-
ited laccase activity in the DAF-stained
Peroxidase as well as laccase activity
were detectable in lignifying tissues of
loblolly pine by histochemical methods
similar to those of Harkin and Obst (4).
The peroxidase activity was present in
both lignifying and nonlignifying tissues
and was readily detected by soluble-en-
zyme assays in all of the crude tissue
extracts studied (Table 3). Cross sections
of woody stems stained more intensely
with DAF and syringaldazine in the pres-
ence of added H202 (Fig. 2D). All tissues
in the cross section, including nonliving
mature xylem, showed peroxidase activity,
in contrast with the more specific staining
of laccase. No staining was obtained, with
or without added H202, in sections that
were boiled before staining.
Our analysis of loblolly pine xylem
laccase challenges the hypothesis (4) that
peroxidase is the only enzyme participat-
ing in lignification'. Laccase is associated
with lignification by four criteria. The
enzyme is found in xylem, is associated
with the cell wall, is present in lignifying
cells, and can oxidize monolignols to
DHP. The role of the laccase purified from
sycamore maple cells in culture is less
certain because the cultured cells do not
make lignin (9, 10), and the relation of
other polyphenol oxidase activities in
stems of sycamore maple (9) to the pine
laccase is not clear. Savidge and Udag-
ama-Randeniya (19) describe a coniferyl
alcohol oxidase activity associated in time
and place with lignification in the cell wall
preparations of differentiating xylem of
conifers. Similarly, in Forsythia sp., an
insoluble cell wall fraction was shown to
catalyze the formation of pinoresinol from
E-coniferyl alcohol with no cofactors oth-
er than oxygen (20). The relation of these
enzyme activities to the pine xylem lac-
case will become clearer once they have
REFERENCES AND NOTES
1. H. Erdtman, Biochem Z. 258, 172 (1933).
2. K. Freudenberg, in Constitution and Biosynthe-
sis of Lignin, K. Freudenberg and A. Neish,
Eds. (Springer-Verlag, New York, 1968), pp.
3. W. Nakamura, J. Biochem. (Tokyo) 62, 54 (1967).
4. J. M. Harkin and J. R. Obst, Science 180, 296
5. T. Higuchi, Wood Sc!. Technol. 24, 23 (1990).
6. R. Goldberg, T. Le, A. Catesson, J. Exp. Bot. 36,
7. A. Imberty, R. Goldberg, A. Catesson, Planta 164,
8. R. Bligny and R. Douce, Biochem. J. 209, 489
9. A. Driouich et al., Plant J. 2, 13 (1992).
10. R. Sterjiades, J. F. D. Dean, K. E. Eriksson, Plant
Physiol. 99, 1162 (1992).
11. A. M. Mayer and E. Harel, Phytochemistry (Ox-
ford) 18, 193 (1979).
12. W. Bao, D. M. O'Malley, R. R. Sederoff, Proc. Natl.
Acad. Sci. U.S.A. 89, 6604 (1992).
13. Glycoprotein was detected by a thymol-sulfuric
acid stain [C. Gerard, Methods Enzymol. 182, 529
14. Cell walls were prepared from 1 kg of differenti-
ating xylem (12). Cell wall proteins were extracted
with 1 M CaCI2, dialyzed, applied to a cation
exchange column (Bio-Rex-70, Bio-Rad, Her-
cules, CA), and eluted with a gradient of NaCI (0
to 2 M; flow rate = 2 ml per minute). Fractions with
laccase activity were concentrated, desalted, and
loaded onto a hydrophobic interaction HPLC col-
umn (Shodex HIC PH-814, Millipore) and then
eluted with a reverse concentration gradient of
ammonium sulfate (2 to 0 M; flow rate = 1 ml per
minute). Active fractions were concentrated, de-
salted, and applied to a reactive dye HPLC col-
umn (Durasphere Cibacron, Alltech, Deerfield, IL)
and then eluted with a gradient of NaCI (0 to 2 M;
flow rate = 1 ml per minute). The specific activity
of the purified enzyme with DAF as the substrate
was 4.6 ,ukatals per milligram of protein (1 ,ukatal =
1 ,umol of product per second). The spectrophoto-
metric laccase activity assay contained 0.68 mM
DAF in 20 mM Bistris buffer (pH 5.9) at 25?C. The
change in absorbance at 600 nm was monitored at
15-s intervals. This assay was reproducible and
linear over time and with different enzyme concen-
15. Laccase was deglycosylated with hydrogen fluo-
ride-pyridine by a method modified from A. J.
Mort and D. T. A. Lamport, Anal. Biochem. 82, 289
16. R. Bereman, unpublished results. The G parallel is
2.076, the G perpendicular is 2.217, and the A
parallel is 74.2 x 10-4 per centimeter [D. J.
Kosman and R. D. Bereman, in Spectroscopy in
Biochemistry, J. Ellis Bell, Ed. (CRC Press, Boca
Raton, FL, 1981), vol. 2, pp. 57-108]. No direct
ESR data support a type 11 copper site, which is
more difficult to detect.
17. The Km values and standard errors were estimat-
ed by nonlinear regression as in R. J. Leatherbar-
row, Enzfitter (Elsevier, Amsterdam, 1987).
18. G. Aulin-Erdtman and R. Sanden, Acta Chem.
Scand. 22, 1187 (1968).
19. R. Savidge and P. Udagama-Randeniya, Phy-
tochemistry (Oxford) 31, 2959 (1992).
20. L. B. Davin, D. L. Bedgar, T. Katayama, N. G.
Lewis, ibid., p. 3869.
21. We thank C.-L. Chen, J. Dean, and E. Wheeler
for their interest and helpful suggestions; J.
Ralph and S. Quideau of the United States
Department of Agriculture (USDA) Dairy Forage
Research Center for substrates; C.-L. Chen for
degraded lignin standards; A; Stomp and T.
LaPasha for assistance with microscopy and
sectioning; R. Bereman for the ESR analysis; C.
S. Levings ll, G. Ward, and K. Korth for assis-
tance with the oxygen electrode measurements;
and M. Gold, N. Lewis, and J. Dean for sharing
results before publication. Supported by North
Carolina State University Forest Biotechnology
Industrial Associates, USDA Forest Service Co-
operative Agreement 29-897, Department of En-
ergy Division of Energy Biosciences (DE-FG05-
92ER20085), and the USDA Competitive Grant
Program (90-37290-5682, 91-37103-6538, and
2 October 1992; accepted 16 February 1993
SCIENCE * VOL. 260 * 30 APRIL 1993