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LIBRARY

Michigan State
University

 

This is to certify that the

thesis entitled

Factors affecting the response of yellow
nutsedge (Cyperus esculentus L.) to 2-chloro-
'2',6'-diethyl-flf(methoxymethyl) acetanilide
(alachlor)

presented by

Thomas F. Armstrong

has been accepted towards fulfillment
of the requirements for

Ph.D. degreein Crop and Soil Sci.

 

 

, - I
Major professor I "\

Date Sept. 8, 1972

 

0-7639

ABSTRACT
FACTORS AFFECTING THE RESPONSE or YELLOW NUTSEDGE (gxzsaus

L.) T0 2-CHLORO-2',6'-DIETHYL-Nr
(METHOXYMETHYL) ACETANILIDE (ALACHLOR)

By
Thomas F. Armstrong

Preplant or preemergence applications of Z-chloro-
2',6'-diethyl-flr(methoxymethyl) acetanilide (alachlor)
at 3.36 or “.48 kg/ha effectively controlled yellow
nutsedge (gypgrug esgulgntus L.) in field and greenhouse
studies. Depth of tubers in the soil did not affect
nutsedge response to alachlor. Alachlor at 3.36 kg/ha
did not effectively control yellow nutsedge in soils with
more than 6% organic matter. Alachlor at 3.7 x 10"6 M
and 18.5 x 10-5 m inhibited growth and killed newly emerging
shoots in Petri dish cultures. However, alachlor did
not inhibit sprouting of yellow nutsedge tubers.

The main site of uptake of alachlor by small yellow
nutsedge plants is the tissue above the tuber. Primarily
acropetal and some basipetal 1”’C-tzranslocation occurred
from 1l‘c-alachlor applications to small seedlings with
2 to 4 cm shoots. Alachlor is metabolized rapidly in yellow
nutsedge to at least one water-soluble metabolite.
Applications of 1tic-alachlor to 15 cm shoots resulted

in limited acropetal 1l‘C-translocation.

FACTORS AFFECTING THE RESPONSE OF YELLOW NUTSEDGE (EIEEBQS
ESQHLEHIHS L.) TO 2-CRLORO-2',6'-DIETRrL-Nr
(METHOXYMETHYL) ACETANILIDE (ALACHLOR)

BY

Thomas Fred Armstrong

A THESIS

Submitted to
Michigan State university
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Crap and Soil Sciences

1972

ACKNONEEDOEMENTS _,

 

 

 

The author wishes to gratefully acknowledge the assistance
of Dr. William Meggitt in conducting this study and in making
my educational experience at M.S.U. possible and rewarding.
A special acknowledgement is extended to Dr. Donald Penner for
his laboratory guidance. The assistance of Drs. E. C. Rossman,
S. N. Stephenson and J. M. TiedJe is also gratefully acknowledged.
Last, but certainly not least, the author would like to thank
his family, Sue, Chris, Cathy and Matt (above),for the sacrifices
they have made to make this day possible.

6
.,1
, ,22

La

TABLE OF CONTENTS

ACKNOWLEDGEMENTS . . . . . .
LIST OF TABLES . . . . . . .
LIST OF FIGURES . . . . . .
INTRODUCTION . . . . . . . .
CHAPTER 1:

LITERATURE REVIEW

0

Yellow Nutsedge (Cyperus esculentus

L.

Biological Control of Yellow Nutsedge

Mechanical Control of Yellow Nutsedge

Chemical Control of Yellow Nutsedge .

Alachlor and Yellow Nutsedge

CHAPTER 2:
ALACHLOR . O O C O C O O 0

Abstract . . . . . . .
Introduction . . . . .
Materials and Methods .
Results and Discussion
Literature Cited . . .
CHAPTER 3:

Abstract . . . . . . .

GROWTH SUPPRESSION

OF YELLOW

NUTSEDGE WITH

THE ABSORPTION, TRANSLOCATION,
OP ALACHLOR BY YELLOW NUTSEDGE

Page
iii
vi

viii

\OUM'lrmNH

11
.11
13
17
37

METABOLISM

IntrOduCtion O O O O I O O O O O O O O O O O O O C

Materials and Methods .

iv

39
39
39
#2

Table of Contents (Cont.)

Results and Discussion .

Literature Cited

CHAPTER 4:

LIST OF REFERENCES

APPENDICES

SUMMARY AND CONCLUSIONS

51

65
68
76

LIST OF TABLES

Table

CHAPTER 2

1.

Yellow nutsedge control 4 weeks after treatment
from preplant incorporated herbicide treatments
on a sandy loam soil with 2.8% organic matter . .

2. Yellow nutsedge control 4 weeks after treatment
from preemergence herbicide treatments on a sandy
loam soil with 2.85 organic matter . . . . . . .

3. Nutsedge response to alachlor 6 weeks after
treatment as affected by depth of tubers in soil

4. Ryegrass bioassay of alachlor leaching over a
6-week-period in greenhouse soil . . . . . . . .

5. Response of yellow nutsedge to preemergence
alachlor and various levels of organic matter
8 weeks following treatment . . . . . . . . . . .

6. Effect of alachlor on nutsedge tuber sprouting
and shoot growth 5 days after treatment . . . . .

7. Effect of tuber exposure time to 3.7 x 10‘6 M
alachlor on nutsedge shoot length . . . . . . . .

8. New shoot emergence and growth of yellow nutsedge
from sand cultures cgntaining sprouted tubers
exposed. to 307 x 10- M 3180,1101“ o o o o o o o 0

CHAPTER 3

l. Site of alachlor uptake by yellow nutsedge . . .

2.

application . . . . . . . . . . .

C-alachlor

Old yellow nutsedge 2 days after

The partitioning and distributionlgf lliC in 10-day

vi

Page

18

19

21

22

24

30

33

34

53

62

List of Tables (Cont.)
Table Figs

3. The separation b TLC of 1"‘c-extracts from I acts
of ll-day-old ye low nutsedge 5 days after c-
alaehlor treatment . . . . . . . . . . . . . . ... 63

V11

LIST OF FIGURES

Figure Page
CHAPTER 2

1. Response of yellow nutsedge to alachlor and various
levels of organic matter weeks following treat- 25
mentoooocooceooooooooooosoooo‘

2. Response of yellow nutse e tO alachlor and various
levels of organic matter weeks following treat- 27
montoooooooooooocoooocooooooo

3. Effect of alachlor on nutsedge shoot growth 5 days
artartrfiatmentoooooossoccoococoa.31

4. New shoot emergence and growth of yellow nutsedge
from sand cultures cgntaining sprouted tubers
exposed to 3. 7 x 10‘ M alachlor. C e no treatment,

A - alachlor for 1 week, and B - alachlor for 4
weeksooocooooo.sooooooooccoco.35

CHAPTER 3

l. Charcoal barrier method that separates alachlor
treated soil from untreated soil . . . . . . . . . . 43

2. Diagram Of charcoal barrier method showing shoot
or root alachlor uptake . . . . . . . . . . . . . . . 45

3. Sites of application of 14C-alachlor to yellow 48
nutsedge p ants. . . . . . . . . . . . . . . . . .

4. Site of alachlor uptake by yellow nutsedge. C a
control, LAl and LA3 a alachlor applied above the
tuber at 1.12 and 3. 36 kg/ha respectively. LBl and
LB3 a alachlor applied below the tuber at 1.12 and
3.36kg/harespeot1vely.o............. 5“

5. Translocation of ltic-alachlor in 15 cm yellow nutsedge
plants. Left: nutsedge plant treate “with “C-
alachlor. Right: radioautograph Of “C-alfichlor
treated plant. Rectangular area is where c-alachlor
was applied . . . . . . . . . . . . . . . . . . . 56

viii

List of Figures (Cont.)

Figure Page

6.

Sites of 1“C-alachlor application on yellow

nutsedge seedlings. A - alachlor application

on small shoot above growing point. B - alachlor
applied on tuber rhizome. Pencil indicates

location of growing point. C a root application
OralaChloroooooooooooooooooooo58

Translocation of inc-alachlor from three sites of
application. Top - nutsedge treated plants and
bottom = radioautograph of treated plants. A, B and
I4. shoot, tuber rhizomes and root application of

c-alachlor'...................60

ix

INTRODUCTION

Man has identified, propagated and used many plants.
Some of these plants are pests in lawns, gardens and
agricultural fields; one of these plants, yellow nutsedge
(Qypgzus esgulentug L.) has become a serious weed problem.
Due to the severity of the nutsedge problem, an economical
control program is needed. Therefore, when a new herbicide
is developed, it should be applied to nutsedge to determine
any toxic effects and deve10p control methods.

A new herbicide, 2-chloro-2',6'-diethyl-nr(methoxymethyl)
acetanilide (alachlor), for use in corn (Z:§.E§X§.L-) and
soybeans [glycine M (L.) Merr.) was reported to be
phytotoxic to yellow nutsedge (l, 10, 13, 22, 24). The
purpose of this study was to more fully understand the
factors affecting the efficacy of alachlor on yellow

nutsedge in field, laboratory, and greenhouse studies.

CHAPTER 1
Yellow nutsedge (Qypgzpfi esgulentpg In)

Both purple (Qyperug potundus L.) and yellow nutsedge
(Qyperus esculentus L.) cause important weed problems in
crops (47, 48, 50, 86). Purple nutsedge has been ranked
as the most important Single weed in the developing
countries of the world (47, 48). Yellow nutsedge is found
in most states of the United States (65, 85, 86), and its
infestation into cropland is increasing (25, 60, 85).

Yellow nutsedge is a perennial sedge with numerous
slender rhizomes ending in tubers or shoots (36). The
morphological characteristics of the plant are affected
by the length of the photoperiod (9, 17, 55). Maximum
shoot and tuber formation from indeterminate rhizome tips
occurs with photOperiods of 16 hr and 8 to 12 hr (55).
Flowering occurs between photoperiods of 12 to 14 hr.
Tubers are also produced by mature plants under long day
conditions (9).

In yellow nutsedge tubers, the buds are located at
the apical end, arranged in sets of three, and positioned
as at points of an equilateral triangle (13). Most tubers
contain two sets of three buds, the second set is inside
the triangle formed by the first set (13). The largest bud
of the outer set sprouts first, and the destiny of sprouted

buds determines the number of buds which ultimately sprout

3

from a given tuber (13). The vascular tissue of rhizomes,
which terminate in tubers, extends through the tubers and
into the buds, roots, and rudimentary leaves of the new
shoot (13).

Both viable seeds and tubers are produced by yellow
nutsedge (9, 10, 45, 84). In one growing season, a single
seedling developed into a stand of plants that produced
a yield of 90,000 seeds of which 51 percent germinated (45).
One tuber in the field produced 1900 plants plus 6900 tubers
in an area 3.2 m2 in 1 year (84). Eight-five percent of
the tubers are found in the top 15.2 cm of soil. Each
plant produces seven to nine tubers, borne singly, on
terminal rhizomes. A tuber produces up to seven shoots,
but a given shoot produces many rhizomes that give rise
to new shoots or tubers (84). These tubers have the capacity
for at least three repeated sproutings (80). Over 605 of
the tuber dry weight, carbohydrate, oil, starch, and protein
are consumed during the first sprouting, but less than 10
percent of these materials are utilized during each of the
remaining two sproutings. Tubers, may be significantly
different in weight, but have the same sprouting percentage (80).

Tuber sprouting is affected by depth of planting in
the soil, soil compaction, dormancy, and mechanical disturbance.
In the soil, tubers sprout readily at 7.6 to 15.2 cm depths;
below this soil level, and in compacted soils sprouting is

reduced (17). Even though sprouting is reduced at lower soil

ll

depths, shoots will grow from tubers 30.5 cm deep in the
field and tubers 50.8 cm to 81.3 cm deep in the greenhouse (84).
In newly formed tubers, the percent sprouting is very low (9,
17, 82). This dormancy can be broken with low temperatures,
chemicals, or by washing in 13 C water (9, 17, 58, 84).
Both mowing and disk-harrow cultivation of a yellow nutsedge
stand significantly increased tuber sprouting during a
period when sprouting from tubers in an undisturbed stand
was low (82). Tillage Operations that exposed tubers on
the soil surface for 2 days lowered sprouting percentage
80 percent (84). Tubers that were exposed to 4 C had a
lower survival than tubers exposed to 22 C. Duration of
dessication did not influence tuber survival, except at
the 4 C temperature (83).

Yellow nutsedge is a persistent pest in many of our
agricultural crops in the United States and the world.

Its infestation into agricultural areas is inereasing (85).

WMQWW
The insect, Eggtza,y§zutan§ Zeller (Olethreutidae), has

been Observed to attack and destroy yellow nutsedge plants

in California. Specific plant injury symptoms were dessication
of the intermost leaves and severe injury to growing points.
This insect is limited in its usefulness as a biological
control agent because of spring emergence of yellow nutsedge
and late summer occurrence of figgtra venutgna Zeller, the

prolific vegetative nature of yellow nutsedge, and insect

parasites on Egctra verutgpg Zeller (62).

5
Ms:hanissl.ssntzsl.2f.xsllsl;nnissdss

In an attempt to eradicate yellow nutsedge, workers
at Cornell university kept nutsedge infested land fallow
for 4 years (9). After four years, 119 tubers/m2 were
found in the top 15 cm of soil compared to 5821 tubers/m2
in the check. These tubers sprouted and eventually new
tubers were formed. Therefore, mechanical methods reduced
the severity of the nutsedge problem but did not eliminate
it.

chemical.ssnirsl.2f.zsllsn.nntssdss

Chemical control offers a means of controlling yellow
nutsedge. Various herbicides have been applied to yellow
nutsedge, but few effectively control this sedge.

Arsenical herbicides have been applied to yellow nutsedge
in turf. Tubers collected from plants treated with 3.36
kg/ha of disodium methanearsonate (DSMA) or monceodium
methanearsonate (MSMA) sprouted in significantly reduced
numbers compared to control tubers (61). small tubers
contained a significantly higher concentration of arsenic
(23 to 33 ppm) than large tubers (4 to 12 ppm) and the
herbicides reduced the vitality of small tubers but had
little or no effect on vitality of large tubers (61).

DSMA and MSMA have provided 80$ or greater control of
yellow nutsedge (28, 59), but MSMA caused injury to the turf (28).

6
In New YOrk State, sodium 2,2-dichloropropionate
(dalapon) applied to yellow nutsedge plants 10.2 on in
height gave complete control in two of four locations (77).

ltic-dalapon indicated there was negible

Investigations using
basipetal movement of 140 after leaf absorption, but there
was excellent distribution after root absorption (77).

In the previous Inc-dalapon experiments only small amounts
of 14
and dalapon had little effect on the viability of intact
tubers (22).

In potatoes treated with 5.6 kg/ha of dalapon in l

c accumulated in the parent tuber of the plant (77).

and 2 applications,the dry weight of yellow nutsedge was
reduced considerably, however the nutsedge was not dead (6).
The highest yield of potatoes was obtained in the aforementioned
experiment with 28 kg/ha of the sodium salt of trichlorOacetic
acid (NaTCA) which gave some control of yellow nutsedge.

Yellow nutsedge plants treated with 3-amino-l,2,4-
triazcle (amitrole) produced tubers containing amitrole.
These tubers rarely sprouted and the seeds had lower germination
(9). Badioautographs of yellow nutsedge treated with lquamitrole,
indicated translocation from the point of application to the
seed (45). Both acropetal and basipetal translocation was
observed in 30.5 cm high yellow nutsedge plants. Amitrole
applied to the foliage translocated to the intact tubers

and greatly reduced their viability (22).

7

EPTC (firsthyl diprOpylthiocarbamate) has been used
to control yellow nutsedge in corn and potatoes (34, 52,
74). Other thiocarbamates, butylate (firsthyl diisobutyi-
thiocarbamate) and vernolate (firpropyl diprOpylthiocarbemate)
have been used in corn and soybeans, respectively (34, 87).
Excellent yellow nutsedge control was obtained with 4.5
to 6.7.kg/ha of EPTC and butylate (29, 34); these two
herbicides provided greater yellow nutsedge control if
applied during or after germination (23). In Illinois,
vernolate equalled EPTC for yellow nutsedge control and
resulted in less injury to soybeans (87).

Yellow nutsedge tubers germinated readily in soil
treated with EPTC. However, shoot growth was suppressed
and emergence delayed, the delay depended on the rate of
application (49). The placement of EPTC in the soil affected
the control of yellow nutsedge (43). As the depth of EPTC
placement in the soil increased, yellow nutsedge control
decreased (44). Butylate placed in the tuber zone effectively
suppressed shoot growth and tuber sprouting, but it was
ineffective when incorporated above or below the tubers (53).
Apparently the thiocarbamates need to be placed near the
yellow nutsedge tubers to give control.

Atrazine [2-chloro-4-(ethylamino)-6-(isoprcpylamino)-
g-triazine] has been applied in a preplant, preemergence

and post-emergence manner to yellow nutsedge (21, 23, 29. 3“. 73).

ll

8

Preplant incorporated applications of atrazine at 2.2 to
4.5 kg/ha applied during or after tuber sprouting controlled
nutsedge (23). Two cultivations following preemergence
applications of atrazine during a dry season significantly
reduced yellow nutsedge stands compared to the use of atrazine
with cultivation (63). Preemergence applications of atrazine
require more rainfall for activation (34). The addition of
non-phytotoxic oil to early postemergence applications of
atrazine increased nutsedge control (21, 23).

Atrazine injured new plants from germinating tubers
but did not kill dormant tubers (9). Translocation studies
with ltic-atrazine, showed that atrazine does not accumulate
in the tubers but is readily translocated throughout the
nutsedge plant (8). This explains why incorporated
applications of atrazine were less effective when tubers
were dormant than at later stages (23).

2,4-D (2,4-dichlor0phenoxy aCetic acid) has been used
for nutsedge control with varying degrees of success.
Bhan et,al. (15) reported that 2,4-D significantly reduced
shoot weight, rhizome weight, and rhizome number 12 days
after application (15). In turf, repeated application of
2,4-D significantly reduced the number of yellow nutsedge
shoots per square foot (28). Other researchers have found
that 2,4-D did not control this sedge (6, 53, 69).

various substituted urea herbicides have also been

tried for yellow nutsedge control (7, 34, 78). Monuron

9
3-(prchlorphenyl)-l,l-dimethylurea applied at 11.2 kg/ha
significantly reduced the number of yellow nutsedge plants (6).
Preemergence applications of monuron at 5.6 kg/ha reduced
top growth by as much as 63% and tuber production by 981 (7).
Preemergence or postemergence applications of 3-(3,4- .
dichlorophenyl-l-methoxy-l-methylurea (linuron) at 2.24 kg/ha
did not control yellow nutsedge (34).

Alachlsz.and.xsllen.nsi§sdse

During 1969, a new herbicide 2-chloro-2',6'-diethyl-
Nr(methoxymethyl) acetanilide alachlor registered for weed
control in soybeans {hlyging mg; (L.) Merré} and corn (z:a,mgyfi
L.) (88) was reported to be phytotoxic to yellow nutsedge
(2, 37, 58, 87, 90). Yellow nutsedge control in corn was
obtained with preplant incorporated or preemergence applications
of alachlor (2, 14, 37); and in soybeans 90 percent control
resulted from preplant incorporation treatments (87).

Under dry conditions, preemergence treatments of alachlor
gave only fair control (27, 87), but a shallow incorporation
of not more than 5 cm provided equal or better control (27,
87).

Alachlor placed near yellow nutsedge tubers in soil
reduced shoot growth (53, 87). Worthington (90) has shown
that alachlor delayed the emergence of yellow nutsedge

plants and increased the number of tuber rhizomes that

10
developed from each tuber. Godke reported significant
reduction of tuber production with preemergence applications
of alachlor (37).

The mode of action of alachlor is unknown. Since
alachlor's chemical structure is similar to nrisoproply-Z-
chloroacetanilide (prOpachlor), it may affect protein
synthesis (26, 57). Possibly nitrate reductase is inhibited
by alachlor (21), or the activation of amino acids is
prevented (57). Alachlor was found to have no detrimental
effect on soil microbial activity at recommended field

rates (67).

CHAPTER 2

Growth suppression of Yellow Nutsedge with Alachlor
Abstract

Preplant or preemergence application of 2-chloroé2',
6'~diethyl-nr(methoxymethyl) acetanilide (alachlor) at
‘3.36 or 4.48 kg/ha effectively controlled yellow nutsedge
(prezufiggggulentus L.) in field and greenhouse studies.
Depth of tubers in the soil did not affect nutsedge response
to alachlor. Alachlor at 3.36 kg/ha did not effectively
control yellow nutsedge in soils with more than 6% organic
matter. Alachlor at 3.7 x lo"6 M and 18.5 x 10'5 M inhibited
growth and killed newly emerging shoots in Petri dish
cultures. However, alachlor did not inhibit sprouting of

yellow nutsedge tubers.
Introduction

Yellow nutsedge is increasing as a weed problem in
most states of the United States (7, 13, 14, 18). Both
viable seeds and tubers are produced by yellow nutsedge (2,
3, 10, 17), but propagation by tubers is the most important
means of dissemination in cultivated crops (2). One tuber
in a Minnesota field produced 1900 plants plus 6900 tubers

in a 3.2 1112 area in 1 year (17).

ll

12

Tuber sprouting is affected by tuber depth in soil,
soil compaction, tuber dormancy, and mechanical disturbance.
In the soil, tubers sprout readily at 7.6 to 15.2 on depths;
below this soil level and in compacted soil, sprouting
is reduced (4). The percent sprouting is very low from
newly formed tubers (2, 4, 16). Both mowing and disk-narrow
cultivation of a yellow nutsedge stand significantly increase
tuber sprouting (l6).

Atrazine [2-chlcro-4-(ethylamino)-6-(isopropylamino)-
artriazinéj has been applied in a preplant, preemergence
and postemergence manner for yellow nutsedge control (5,
6, 8, 9, l5). Preplant incorporated applications of atrazine
at 2.2 to 4.5 kg/ha controlled yellow nutsedge if applied
during or after, rather than before tuber sprouting (6).
Preemergence applications of atrazine require rainfall for
activation (9). The addition of non—phytotoxic oil to
early postemergence applications of atrazine increased
nutsedge control (5, 6). Atrazine injured new plants from
germinating tubers but did not kill dormant tubers (2).

In 1969, alachlor was registered for weed control in

soybeans [m m (L.) MerrJand corn (2; m L.); and
reported to be phytotoxic to yellow nutsedge (1, l3, 21).1' 2

 

1Godke, D. L. 1969. The control of yellow nutsedge with
alachlor’and an alachlor plus linuron mixture., M.S. Thesis.
Southern Illinois University, Carbondale. 58p.

2Worthington, J. P. 1971. The effect of alachlor and

Mon-097 on the growth of yellow nutsedge and the uptake of

1l‘C-labeled alachlor by yellow nutsedge and soybeans. Ph.D.
Thesis. Ohio State University, Columbus. 62p.

13

Alachlor placed near yellow nutsedge tubers in soil
reduced shoot growth (11, 20). Worthington3 has shown
that alachlor delayed the emergence of yellow nutsedge
plants and increased the number of tuber rhizomes that
developed from each tuber. Godke“ reported a significant
reduction in tuber production with preemergence applications
of alachlor.

'The objectives of this study were to evaluate the
response of yellow.nutsedge to alachlor in the field,
greenhouse, and laboratory and to deve10p an effective

yellow nutsedge control program with alachlor.
Materials and Methods

£1§1g_§§pgig§ during 1969, 1970 and 1971 were conducted

at E. Lansing, Michigan on a Conover sandy loam (2.8%

organic matter) With a native infestation of yellow nutsedge.

The field was fall plowed, the plots were 3.1 by 10.8 m,

and the treatments were replicated three times in a randomized
complete block design. The preplant treatments were incorporated
twice with a springtooth harrow and Michigan 400 corn was
planted. The herbicides were applied with a tractor mounted
sprayer at 30 psi and 215 L/ha. The plots were not cultivated,

and were evaluated approximately 4 weeks after herbicide

 

3min-
“Codke, D. L. op. cit-

14

application. Yellow nutsedge control was rated on a 0
to 10 scale where O - no control and 10 - complete control
or kill. Only the 2 middle rows (152 cm) of each 4-row
plot were rated.
nggnhgp§§_§tpgi§§_were conducted to determine yellow
nutsedge response to alachlor as affected by depth of
tubers in a loam soil. Tubers that received 6 weeks of
7 C cold treatment were sprouted in Petri dishes. Nine
sprouted tubers each with less than 1 cm shoot length
were planted at either 2.5, 10.2, or 20.3 cm depths in
each 30.5 by 33.0 by 43.2 cm wooden box. water was added as
needed throughout the experiment to maintain soil moisture.
Preemergence applications of alachlor at either 0.00, 2.24
or 4.48 kg/ha were made for each tuber depth. After 6
weeks, the yellow nutsedge was harvested and dry matter
determined. The treatments were replicated three times
in a randomized complete block design.

After harvest of the yellow nutsedge, a perennial
ryegrass bioassay was used to determine alachlor residue
in the loam soil. Soil samples from the previous experiment
of 0.0 to 5.1, 5.1 to 10.2, and 10.2 to 15.2 cm depths were

placed in a 750 cm3 styrofoam container and 30 ryegrass

seeds were planted in each container. After 2 weeks of
growth, the ryegrass was harvested and mg fresh weight/plant

recorded.

15

The response of yellow nutsedge to various levels
of organic matter and preemergence alachlor treatments
was examined. A Ccnover sandy loam sell with 2.84%
organic matter and a muck soil with 81.40% organic matter
were mixed together in various volume ratios to obtain
2.84%, 4.57%, 5.55%, 10.28% and 81.40% organic matter
soils. One tuber was planted 3 cm deep in each 473 ml
container containing the respective level of organic
matter. Preemergence applications of alachlor were made
at either 0.00, 1.12 or 3.36 kg/ha. Shoot emergence and
shoot height were recorded after 8 weeks. water was
added throughout the duration of the experiment to maintain
soil moisture. The treatments were replicated seven times
in a completely randomized design.
Lgpgzgtgzy_§tng1g§ were used to determine the effect of
alachlor on yellow nutsedge tuber sprouting and growth.
Tubers were sprouted in Petri dishes containing solutions
of O, 3.7 x 10'6 M, or 18.5 x 10"5 M alachlor. Fifteen
tubers were placed in each Petri dish. After 5 days the
percent of tubers sprouting and shoot length were recorded.
Each treatment was replicated three times in a completely
randomized design.

The effect of tuber exposure time to 3.7 x 10'"6 M

alachlor on yellow nutsedge shoot length was determined

by placing tubers in Petri dishes containing solutions
of 3.7 x 10'6 M alachlor for o, 8, 24, as, 72, 96, 120 and

16
144 hrs. At the end of each time exposure, the respective
tubers were rinsed off six times with distilled water and
then replaced in Petri dishes, five per dish, containing
only distilled water. Shoot length was recorded at l, 2,

3, and 4 weeks after the start of experiment. Each treatment
was replicated three times in a completely randomized design.
Tubers were sprouted in 0 and 3.7 x 10"6 M alachlor

solutions in Petri dishes for 1 week. The sprouted tubers
were then transfered to 293 ml plastic cups containing ,
washed sand. The tubers were planted 2 cm deep, and number
of shoots present on each tuber and the shoot length were
recorded. One half of the alachlor treated tubers received
Hoaglunds no. 1 nutrient solution with alachlor for 3 weeks.
The cups containing the tubers were placed in a growth chamber
at 25 C. At the end of the experiment new shoot emergence
from the cups and shoot length were recorded. The treatments
were replicated four times in a completely randomized design.
All data presented is the mean of two experiments with two

or more replications per experiment.

Results and Discussion

The amount of rainfall during 10 days after the start
of treatment influenced the degree of yellow nutsedge control.
The amounts of rainfall during the 10 days after herbicide
application for each year were the following: 1969, 3.3 cm;
1970, 1.8 cm and 1971, 1.0 cm. The rainfall in both 1969
and 1970 appeared adequate, whereas the rainfall in 1971
provided less than adequate moisture for weed control.
Preplant incorporated treatments of both atrazine and
alachlor controlled yellow nutsedge (Table 1). In 1971
when the rainfall was less than adequate, the incorporation
of the atrazine at 4.48 kg/ha provided greater yellow nutsedge
control than the preemergence atrazine application. Pre-
emergence application of alachlor at 4.48 kg/ha controlled
more yellow nutsedge than did 2.24 kg/ha (Table 2). This
was especially ture for 1971 when rainfall was less than
adequate. Both preplant and preemergence treatments of
alachlor and atrazine combinations controlled 76 to 88%
of the yellow nutsedge. A minimum rate of 3.36 kg/ha of
alachlor was necessary for yellow nutsedge control. Preplant
treatments with shallow incorporation or preemergence
treatments of alachlor at 3.36 or 4.48 kg/ha controlled
80$ or more of the yellow nutsedge. With less than adequate
rainfall, preplant incorporated treatments of alachlor were more
effective than preemergence treatments; but with adequate

rainfall (1.27 to 2.54 cm rainfall within 10 days after

17

18

Table 1. Yellow nutsedge control 4 weeks after
treatment from preplant incorporated herbicide
treatments on a sandy loam soil with 2.8x organic

 

 

matter.

Chemical Rate Weed control ratinga
(ks/ha) (1969) (1970) (1971) (mean)

Atrazine. 4.48 9.0 9.7 9.7 9.6
Alachlor 3.36 8.3 8.7 9.0 8.7
Atrazine

+ 1.12 + 2.24 7.5 7.0 8.3 7.6
alachlor
Atrazine

+ 1.12 + 4.48 8.2 8.7 8.3 8.4
alachlor
Atrazine

+ 2.24 + 2.24 8.5 8.7 8.3 8.6
alachlor

 

aO-No control; lO-complete control or kill.

19

Table 2. Yellow nutsedge control 4 weeks after
treatment from preemergence herbicide treatments
on a sandy loam soil with 2.8% organic matter.

 

 

Chemical Rate weed control rating‘
(ks/ha) (1969) (1970) (1971) (moan)

Atrazine 4.48 8.8 7.0 5.3 7.0
Alachlor 2.24 7.5 8.7 5.7 7.3
Alachlor 4.48 8.7 8.0 8.3 8.3
Atrazine

+ 2.24 + 2.24 8.5 8.0 7.7 8.1
alachlor
Atrazine

+ 2.24 + 3.36 8.8 9.0 8.7 8.8
alachlor
Atrazine

+ 1.12 + 2.24 8.5 8.3 7.0 7.9
alachlor
Atrazine

+ 1.12 + 3.36 8.7 8.0 8.0 8.2
alachlor

 

aO-No control; lO-complete control or kill.

20
herbicide application), both preemergence or preplant
incorporated alachlor treatments effectively controlled
yellow nutsedge. Postemergence alachlor applications
on yellow nutsedge were not effective.

The depth of the tubers in the soil did not affect
the response of yellow nutsedge to alachlor (Table 3).
Yellow nutsedge shoot dry matter from tubers planted 2.5 cm
which received preemergence alachlor applications was
not significantly different from that of tubers planted
10.2 cm or 20.3 cm deep and received preemergence applications
of alachlor. Both treatments significantly suppressed
nutsedge growth. The three controls were significantly
different in growth from each other because they emerged
at three different times. Shoots emerged first from the
tubers planted 2.5 cm deep then shoots emerged from the 10.2 cm
deep tubers and finally the shoots emerged from tubers
planted 20.3 cm deep.

The ryegrass bioassay for alachlor indicated that
alachlor was present primarily in the top 5 cm of the loam
soil indicating only limited leaching (Table 4).

The response of yellow nutsedge to alachlor was
influenced by the percent organic matter present in the
,soil. The number of yellow nutsedge shoots emerging from
soil with 5.55% organic matter or greater and treated with

a preemergence application of 1.12 kg/ha of alachlor was

21

Table 3. Nutsedge response to alachlor 6 weeks
after treatment as affected by depth of tubers

 

 

in soil.
Tuber depth Alachlor rate Dry mattera
# (om) (kg/ha) M“ (g/mZ)
2.5 0.00 154.6 a
2.24 8.5 de
4.48 10.6 de
10.2 0.00 123.7 b
2.24 6.7 e
4.48 3.2 9
20.3 0.00 68.5 c
2.24 24.1 d
4.48 5.9 e

 

a
Means with common letters are not significantly different
at the 5% level, Duncan's Multiple Range Test.

22

Table 4. Ryegrass bioassay of alachlor leaching
over a 6-week-period in greenhouse soil.

 

 

 

Depth of soil sample Ryegrass growth in 2 wk
filachlor (kgéha)
2.241al 4.48“
(cm) (mg fresh wt/plant)
0.0 to 5.1 14.7 a 12.4 a
5.1 to 10.2 28.0 b ' 24.4 b
10.2 to 15.2 26.1 b 21.7 b

 

aMeans within columns with common letters are not significantly
different at the 5% level by Duncan's Multiple Range Test.

23

similar to the number of shoots emerging in soils not
receiving alachlor (Table 5). The 2.84% and 4.571 organic
matter soils had significantly less shoots emerging when
alachlor was applied. Preemergence application of
alachlor at 1.12 kg/ha regardless of percent organic
matter did not reduce shoot height. Alachlor applied
at 3.36 kg/ha to soils with 5.55% or less organic matter
significantly reduced shoot emergence. Four weeks after
either 1.12 kg/ha or 3.36 kg/ha alachlor application the
shoot height for yellow nutsedge plants were similar,
except in the high organic matter soils (Figure 1).
But after 8 weeks of growth, only the yellow nutsedge
grown in low organic matter soils with less than 5.55%
organic matter had significantly reduced shoot height
(Table 5 and Figure 2).

Alachlor has no significant effect on tuber sprouting,
but it significantly reduced shoot elongation (Table 6).

Concentrations of 3.7 x 10"6 M or 18.5 x 10-5 h alachlor
did not affect the sprouting of the yellow nutsedge tubers.

The low concentration of 3.7 x 10"6 M alachlor dramatically
prevented the growth of yellow nutsedge shoots (Figure 3).
After only 8 hr of treatment, shoot length was significantly
reduced (Table 7).

Fellowing the treatment of tubers with 3.7 x 10'6 n

alachlor for 1 week, more shoots emerged after 4 weeks

24

Table 5. Response of yellow nutsedge to preemergence
alachlor and various levels of organic matter 8 weeks
following treatments.

 

 

Alachlor Organic matter Emergeneea Shoot heigh£:_
(ks/ha) m (shoots/95 cmz) (on)
0.00 2.84 14.9 fg 24.0 c
4.57 13.1 fg 25.9 cd
5.55 12.4 fg 1 24.1 c
10.28 13.4 fg 23.3 bc
81.40 18.1 h 25.8 c
1.12 2.84 7.6 cd 19.2 b
4.57 9.2 de 20.5 be
5.55 12.4 fg 24.9 c
10.28 13.8 fg 22.6 be
81.40 19.1 h 24.9 c
3.36 2.84 1.9 a 5.6 a
4.57 3.6 ab 10.8 a
5-55 5.8 be 17.5 b
10.28 11.3 eg 21.2 be
81.40 9.3 de 20.2 bo

 

aMeans within columns with common letters are not significantly
different at the 5% level, Duncan's Multiple Range Test.

25

Figure 1. Response of yellow nutsedge to alachlor
and various levels of organic matter 4 weeks
following treatment.

26
0.00 MiG/HR

 

w 4: ~ »
2.8 4.6 5.6 l0.3 8L4 °/oO.M.

I.l2 KG_/HH

-aLaLM/

2.8 4.6 5.6 l0.3 8|..4°/oOM.

3.363” H3

---11

2.8 4.6 56 l0.3 8|..4°/oOM

27

Figure 2. Response of yellow nutsedge to alachlor
and various levels of organic matter 8 weeks
following treatment.

28

514 9%0Ji

OQOKG/HR
/, ‘ Iv

. \443'3

," 1 ‘fi.:

5.6 I0.3

2.8 4.3

IJZKG/Hfl

 

4.6 5.5 10.3" 81.49.03.

29
than when alachlor was present for the entire 4 weeks
(Table 8). Apparently alachlor upset apical dominance in
tubers. Shoot length was significantly different among all
three treatments: no alachlor present, alachlor present
for only one week, and alachlor present for four weeks
(Figure 4). The tallest shoots occurred when no alachlor
was present, and minimum shoot growth resulted from 4 weeks
of alachlor treatment. Alachlor did not kill the tubers,
it suppressed shoot growth from these tubers.

Alachlor is an effective herbicide for controlling
yellow nutsedge growth if it is present in the soil during
the time of tuber sprouting. .Yellow nutsedge control with
alachlor is dependent on proper timing and placement of the

herbicide and adequate rainfall after application.

30

Table 6. Effect of alachlor on nutsedge tuber
sprouting and shoot growth 5 days after treatment.

 

 

Alachlor Sproutinga Shoot elongationg
(cone) (5) (mm/plant)

0 94.5 a 15.4 a

3.7 x 10'“5 M 94.3 a 3.3 b

18.5 x 10'5 M 93.3 a 2.6 b

 

aMeans within columns with common letters are not significantly
different at the 5% level, Duncan's Multiple Range Test.

31

Figure 3. Effect of alachlor on nutsedge shoot
growth 5 days after treatment.

 

 

333E393 8.282.

33

Table 7. Effect of tuber exposure time to 3.7 x 10"6 M
alachlor on nutsedge shoot length.

 

Exposure time to

Nutsedge shoot lengtha

 

 

 

alachlor
. 1 wk 2 wk 3 wk_I;fi ‘4 wk
(mm7plant) (nunfplant) (min/plantT (prlant)

(hr)

0 14.5 c 57.8 c 85.5 d 87.5 d

8 9.5 b 25.8 b 38.5 c 37.8 be
24 4.2 a 11.5 ab 15.7 abo 16.3 abc
48 2.5 a 15.0 ab 33.8 bc 42.2 c
72 2.8 a 6.8 a 11.0 ab 12.0 ab
96 2.3 a 7.3 a 14.2 abo 19.2 abc
120 1.3 a 4.3 a 16.3 abc 19.8 abc
144 3.2 a 4.5 a 5.0 a 5.3 a

 

a
Means within columns with common letters are not significantly
different at the 5% level by Duncan's Multiple Range Test.

Table 8.~ New shoot emergence and growth of yellow
nutsedge from sand cultures containing sprouted
tubers exposed to 3.7 x 10"6 M alachlor.

 

 

Treatment New shoot emergencei ‘ Shoot length‘
(number/culture) (cm/shoot)

No treatment 0.9 ab 19.1 a

Alachlor for 1 week 2.1 a 10.0 b

Alachlor for 4 weeks 0.0 b 0.9 c

 

aMeans within columns with common letters are not significantly
different at the 5% level by Duncan's Multiple Range Test.

35

Figure 4. New shoot emergence and growth of yellow
nutsedge from sand cultures containing sprouted
tubers exposed to 3.7 x 10'”6 M alachlor, C - no

treatment, A - alachlor for 1 week, and B . alachlor
for 4 weeks.

1.

2.

3.

5.

7.

8.

9.

10.

11.

Literature Cited

Andrews, 0. N., C. v. Derting, and D. F. Jones. 1971.
An evaluation of alachlor herbicide combinations
in corn. Proc. Southeast. weed Sci. Soc. 24:138-145.

Bell, R. A., w. R. Lachman, E. M. Hahn, and R. D. Sweet.
1962. Life history studies as related to weed
control in the northeast. 'I Nutgrass. Rhode Island
Agr. EXpt. Sta. Bull. 33p.

Bell, R. A. and E. Larssen. 1960. Experiments with
germination of northern nutgrass seed. Proc.
Northeast. weed Contr. Conf. 14:45-48.

Fundy, 0. M., w. F. Donalley, and E. M. Rahn. 1960.
Growth and development of northern nutgrass as
affected by certain environmental conditions.
Proc. Northeast. weed Contr. Conf. 14:44.

Colby, S. R. 1967. Preemergence and postemergence
control of yellow nutsedge with herbicide
combinations and oils. Proc. Northeast. weed
Contr. Confe 21:307’31he

Doty, C. H. and R. D. Sweet. 1972. Yellow nutsedge
control as influenced by time of treatment. Proc.
Northeast. Weed Sci. Soc. 26:137-145.

Duke, w. B. 1972. Nutsedge alert. Agrichemical Age.
p. 22.

Durfee, J. w., w. R. Lachman, and w. C. Lincoln Jr.
1960. Control of northern nutgrass with ep am
and atrazine. Proc. Northeast. weed Contr.
Conf. 1n:214-216e .

Fertig, S. N. and A. H. Furrer, Jr. 1965. Results
of 1964 research on weed control in corn. Proc.
Northeast. Weed Contr. Conf. 19:208-216.

H111, Ea Be, We Be LaChmBn, and De Ne Maynard. 1963s
Reproductive potential of yellow nutsedge by seed.
weeds 11:160-161.

Ingle, E. L. and A. D. Worsham. 1971. Effect of soil
placement of several herbicides on yellow nutsedge
erus esculentus) tubers. Proc. Southeast Heed

5°1e SOCe 2E:TBE-192e

37

12.

13.

14.

15.

16.

17.

18.

19.

20.

38

Keeley, P. E., C. R. Carter, and J. R. Miller. 1972.
Evaluation of the relative phytotoxioity of
Egrgiciges to cotton and nutsedge. weed Sci.

: -7e

Keeley, P. E. and R. J. Thullen. 1972. Nutsedge
control. Agrichemical Age. p. 8.

Lewis, W. M. and A. D. Worsham. 1970. The ten worst
weedz-og field craps. Nutsedge. Crops and Soils
22:1 1 .

Palmer, R. D. and W. K. Porter, Jr. 1959. The metabolism
of nutgrass (szg:n§,22§ungu§ L.) I. The influence
of various oxygen and carbon dioxide levels upon
germination and respiration. Weeds 7:481-489.

Taylorson, R. B. 1967. Seasonal variation in sprouting
and available carbohydrate in yellow nutsedge
tubers. Weeds 15:22-24.

Tumblesone M. E. and T. Kommedahl. 1961. Reproductive
i

poten al of gypgzn§,g§gylgn§g§ by tubers.

U.S.D.A. 1968. Extent and cost of weed control with
herbicides and an evaluation of important weeds,
1965. Agr. Res. Ser. 85pp.

U.S.D.A. 1970. Selected weeds of the United States.
Agr. Handbook No. 366. Agr. Res. Ser.

wax. LO MO. E0 We StOlleI‘, R. W. Slife’ 8111 R. N.
Andersen. 1972. Yellow nutsedge control in
soybeans. Weed Sci. 20:194-201.

CHAPTER 3

The Absorption, Translocation, and Metabolism of
Alachlor by Yellow Nutsedge

Abstract

The main site of uptake of 2-chloro-2',6'-diethyl-N-
(methoxymethyl) acetanilide (alachlor) by small yellow
nutsedge (gypgzufi eggulentus L.) plants is the tissue above
the tuber. Primarily acropetal and some basipetal 140

translocation occurred from 14C-alachlor applications to

small seedlings with 2 to 4 cm shoots. Alachlor is
metabolized rapidly in yellow nutsedge to at least one
water-soluble metabolite. Applications of 1l‘C-alachlor

to 15 cm shoots resulted in limited acropetal 140 trans-

location.
Introduction

The perennial, yellow nutsedge, is a serious weed
problem in the United States and many other parts of the
world (2, 3, 9, 10).

Yellow nutsedge has numerous slender rhizomes
ending in tubers or shoots (l). Tubers in the soil will
produce tuber rhizomes which may form basal bulbs below

the soil surface (5. 8). Shoots, rhizomes, and roots

39

40
are produced from a basal bulb (5, 8). Thus, there are
many potential sites for alachlor uptake by yellow nutsedge
to explain growth inhibition (4, 12). Alachlor also delays
nutsedge emergence and reduces tuber production.1'2

Alachlor uptake has been reported to be greater in the
susceptible species, barley, than tolerant corn.3 Alachlor
was absorbed by the roots and then translocated in the xylem
to the tips of the older leaves. In barley, more alachlor
was present in the shoot than.in the root following treatment.
Thin layer chromatography (TLC) of the water-soluble extracts
from barley shoots and roots showed that most of the 1”c
was found in one major metabolite and one minor metabolite
occurring in each shoot and root.

Similarly, 2-chloro-N-iSOprOpylacetanilide (prOpachlor)
was metabolized to water-soluble metabolites in corn and barley
(7). Both resistant and susceptible species have the ability
to metabolize chloroacetamides (6). The basis for selectivity

in the case of chloroacetamide herbicides may be the ability

1Godke, D. L. 1969. The control of yellow nutsedge
with alachlor and an alachlor plus linuron mixture. M. S.
Thesis. Southern Illinois University, Carbondale. 58p.

2Worthington, J. P. 1971. The effect of alachlor'and
M N-097 on the growth of yellow nutsedge and the uptake of
1 C-labeled alachlor by yellow nutsedge and soybeans. Ph.D.
Thesis. Ohio State University, Columbus. 62p.

3Hamill, A. s. 1971. Bases for the interaction of
alachlor, butylate or chlorbromuron with carbofuran on barley
in? corn. Ph.D. Thesis. Michigan State University, E. Lansing.
2 p.

 

41
of resistant plants to metabolize them at a rate sufficient
to keep their levels below that required for growth inhibition (6).
The objective of this study was to determine the site
of uptake, the translocation, and the metabolism of alachlor
in order to explain growth inhibition in young yellow
nutsedge plants.

Materials and Methods

The site of alachlor uptake was determined by applying
alachlor separatly to yellow nutsedge roots or shoots.
An activated charcoal layer was used to isolate the root
and shoot regions to prevent herbicidal movement (Figure l
and 2). Alachlor was applied at 0, 1.12 or 3.36 kg/ha
above or below the charcoal barrier. water was surface
applied throughout the duration of the experiment to '
maintain soil moisture for nutsedge growth. Shoot emergence
and height were recorded at both 2 and 4 weeks after the
start of the treatment. The values reported in Table 1
are the means of two experiments with ten replications
per experiment.

14

Uniformly ring-labeled C-alachlor was applied to

a 15 cm tall yellow nutsedge plant to study herbicidal

14C-alachlor had a specific activity

translocation. The

of 1.02 no per u mole. Each plant received a 5 n1 drop

containing 0.25 no placed on a mature leaf. After 2 days

of treatment, the plants were freeze dried and radioautographed.
The translocation of 14c-aiaehior was studied in small

yellow nutsedge seedlings with shoots 2 to 4 cm in length

above soil surface by applying 5.u1 with 0.25 no of 1"‘C-alachlor

to one of three different sites of application: on the shoot

above the growing point, on the tuber rhizome below the

growing point or on the roots (Figure 3).

42

43

Figure l. Charcoal barrier method that separates
alachlor treated soil from untreated soil.

5011 Surface

iIl2IKI72I5IIIIQIKIVEIKAFEIEQFEIKIVQIKII'
Tuber 2.0 cm

IIIII‘[Illlllllllllllllllllllliéfilllllllllllllll llllllllllllllll

    
  
   

  

 

Charcoal Barrier

45

Figure 2. Diagram of charcoal barrier method
showing shoot or root alachlor uptake.

 

46

Soil Surface

Jll’mnIl’

Shoot
Uptake

Illllfllllllllllllllllllll lllllllHllllllllllllllflflH .E’

h

Root

Uptake

Charcoal Barrier

47
After 2 days of treatment the experiment was terminated and

the plants radioautographed. The plants were then homogenized

and extracted for 14

u

C-alachlor and metabolites by method of

1L‘C-labeled

Hamill. The samples were extracted for
materials for 6 hr with 10 m1 of 80 percent acetone.

To facilitate extraction the vials containing the samples
were placed in a reciprocating shaker to agitate the sample.
Following extraction the vials were centrifuged at 455 x g
for 5 min. The supernatant fluid was removed, stored, and
the pellet extracted as before for 8 hr, centrifuged, and
the corresponding supernatants fluids combined. The super-
natant volume was reduced under nitrogen at 30 C. After

a third extraction with 10 m1 of 100 percent acetone for

10 hr, the homogenate was filtered through No. l Whatman
filter paper. The filtrate was added to the previously
obtained supernatant fluid. The volume of the combined
filtrate-supernatant was reduced under nitrogen to 4 ml.

One ml of hexane was added to the extract with a Vortex
test tube stirrer. The mixture was centrifuged at 455 x g
for 10 min, the vials sealed, and then placed in a freezer
for 1 hr to facilitate separation of the hexane and aqueous
fractions. Both the hexane and aqueous fractions were

assayed for radioactivity with a scintillation spectrometer.

“mi.

 

48

Figure 3. Sites of application of 1"‘C-alachlor
to yellow nutsedge plants.

 

49

SHOOT

 

__ SOIL SURFACE

_1 TUBER RHIZOME

1 ROOTS

 

50
A portion of the dried acetone-insoluble residue
was weighed and combusted for radioassay by the Schoeniger
combustion method of Wang and Willis (13).

ll‘C-alachlor

Metabolism was studied by applying
to 4 to 6 cm shoots as previously described. After 5
days of treatment, the plants were homogenized in 20 m1
of 80 percent acetone with a Waring blender for 2 min and
filtered through No. l Whatman filter paper. The filtrate
was reduced to a 2.0 m1 aqueous fraction. Two m1 of hexane
was added to the filtrate, mixed, and then put in a freezer
over night to facilitate separation of the two fractions.
The volume of each fraction was reduced to 0.1 ml and 50 ul
of each were spotted on 250 micron thick silica gel H thin
layer plates. The plates were developed in either a polar
or a non polar solvent system. The nonpolar solvent system
was petroleum ether:chloroform:95 percent ethanol (7:2:1;
v/v/v); and the polar solvent system was l-butanol:g1acial
acetic acidéwater (12:3:5: v/v/v). After development for
15 cm on the plate, the plates were divided into 1 cm bands
and the radioactivity determined as previously described.

Results and Discussion

Reduction in yellow nutsedge shoot growth occurred
when alachlor was absorbed from the soil by the tissue above
the tuber (Table 1). Both emergence of shoots and shoot
height were affected. The absorption of alachlor by roots
did not reduce plant growth (Figure 4).

Foliar application of ll‘C-alachlor resulted in limited
acropetal and no basipetal translocation of alachlor in
15 cm tall plants (Figure 5). Postemergence applications
of alachlor probably do not control yellow nutsedge because
alachlor is not translocated to the growing point.

When ll”C-alachlor was applied on the shoot, tuber
rhizome, or root of small nutsedge seedlings (Figure 6)
differences in translocation occurred (Figure 7). The
14C-material translocated from the shoot and tuber rhizome

to the growing point, but the majority of the 1“C from

1M'C-alachlor applied to the root did not move. Alachlor

was absorbed by the shoots of very young plants, and was
translocated acropetally or basipetally to the yellow
nutsedge growing points killing the plants.
Two days after ll‘C-alachlor application, 81 percent
Of the lac-labeled material was metabolized by yellow
nutsedge to a water-soluble product (Tables 2 and 3).
Only one major water-soluble metabolite of alachlor

was present in the water-soluble fraction after 5 days.

51

52

This is similar to Hamill's results showing rapid alachlor
metabolism to a water-soluble product in barley, a susceptible
species.

Alachlor absorption by small nutsedge seedlings through
the shoot or tuber rhizome and subsequent translocation
to the growing point appears responsible for reduced emergence
of shoots, shoot height, and eventual death of the young
plant.

53

Table 1. Site of alachlor uptake by yellow

 

 

nutsedge.
Alachlor Location of Shoot emergencea Shoot hei hfi
rate herbicide 2 Wk 4 Wk 2W
(ks/ha) (z) (1) (cm/plant)
0 - 75 o 95 c 3.85 b 10.4 b
1.12 above barrier 30 b 50 b 0.22 a 1.1 a
3.36 above barrier 0 a 15 a 0.00 a 0.0 a
1.12 below barrier 95 d 100 c 5.01 c 9.8 b
3.36 below barrier 95 d 95 c 5.32 c 11.4 b

 

a
Means within columns with common letters are not significantly
different at the 5% level by Duncan's Multiple Range Test.

54

Figure 4. Site of alachlor uptake by yellow nutsedge
C 8 control, LAl and LA3 - alachlor applied above the
tuber at 1.12 and 3.36 kg/ha respectively, LBl and
LB3 - alachlor applied below the tuber at 1.12 and
3.36 kg/ha respectively.

55

 

56

Figure 5. Translocation of 14C-alachlor in 15 cm

ye low nutsedge plants. Left: nutsedge plant Ereated

with uc-alachlor. Right: radioautograph of 1 C-

akachlor treated plant. Rectagular area is where
C-alachlor was applied.

57

 

58

Fi ure 6. Sites of 1"‘C-alachlor application on
ye low nutsedge seedlings. A - alachlor applied
on small shoot above growing point. B . alachlor
applied on tuber rhizome. Pencil indicates
location of growing point. C 3 root application
of alachlor.

59

 

 

6O

Fi ure 7. Translocation of 14C-alachlor from three

si es of application. Top = nutsedge treated plants
and Bottom = radioautographs of treated plants.

A, B and C = shoot, tuber rhizome and root application
of uC-alachlor.

61

Ill will: Ill I

 

62

Table 2. The partitioning and distribution of 11+c

in 10-day-old yellow nutsedge 2 days after 140-
alachlor application.

 

Treatment Acetone- water- Hexane-
insolublea solublea solublea

 

(dpm/mg) (arm/ms) (dpm/ms)

Shoot
Above growing point 1426 b 25461 b 6321 b
Growing point 790 a 2040 a 390 a
Below growing point 13 a 117 a 26 a
Tuber rhizome
Above 14C spot 33? b 2796 a 164 b
Below 1"’C spot 11 a 17 a 20 a
Root
Above 140 spot 94 b 451 a 70 b
Below 14C spot 15 a 53 a 22 a

 

a
Means within treatment columns with common letters are not
significantly different at the 5% level by Duncan's Multiple
RangerTest.

63

Table 3. The separation by TLC of 1L’C-extracts from
shoots of ll-day-old yellow nutsedge 5 days after

C-alachlor treatment.a

 

Percent of total spotted

 

 

Treatment Rf 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0
Water-soluble 81 3b

fraction

Hexane-soluble 2 14b

fraction

 

3TLC plates were developed in petroleum ether:chloroform:95
percent ethanol (7:2:1; v/v/v).

bThe 11+C-alachlor standard co-chromatographed at this Rf.

1.

7.

8.

9.

10.

11.

12.

Literature Cited

Gleason, H. A. and A. Cronquist. 1968. Manual of
vascular Plants. D. van Nostrand Company, Inc.,
Princeton, New Jersey. 810p.

Holm, L. 1969. Weeds problems in develOping countries.
Weed SCie 17:113-118e

HOlt, Ee Ce, Je Le Fanion, We We Allen, and Ge Ge
McBee. 1967. Arsenic translocation of nutsedge
tuber systems and its effect on tuber viability.
Weeds 15:13-15.

Ingle, E. L. and A. D. Worsham. 1971. Effect of soil
placement of several herbicides on yellow nutsedge
(Cypgzua ) tubers. Proc. Southeast
Weed Sci. Soc. 2 :188-192.

Jansen, L. L. 1971. Morphology and photoperiodic
responses of yellow nutsedge. weed Sci. 19:210-219.

Jaworski, E. G. 1969. Analysis of the mode of action
of herbicidal cflbchloroacetamides. J. Agr. Food
Chem. 17:165-170.

Lamoureux, G. L., L. E. Stafford, and F. S. Fanaka.
1971. Metabolism of 2-chloro-N-isopropylacetanilide
(propachlor) in the leaves of corn, sorghum,
sugarcane, and barley. J. Agr. Food Chem. 19:346-350.

Stoller, E. W., D. P. Nema, and V. M. Bhan. 1972.
Yellow nutsedge tuber germination and seedling
development. Weed Sci. 20:93-97.

U.S.D.A. 1968. Extent and cost of weed control with
herbicides and an evaluation of important weeds,
1965. Agr. Res. Ser. 85p.

U.S.D.A. 1970. Selected weeds of the United States.
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wang, C. H. and D. L. Willis. 1965. Radiotracer
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wax. Le Me. Be We StOller, Be We Slife, and Be Ne

Andersen. 1972. Yellow nutsedge control in
soybeans. Weed Sci. 20:194-201.

64

CHAPTER 4
SUMMARY AND CONCLUSIONS

Field, greenhouse and laboratory studies were conducted

to evaluate the response of yellow nutsedge to alachlor,

and determine the site of uptake, the translocation and

the metabolism of alachlor by yellow nutsedge.

The results of these investigations are summarized

as follows:

1.

2.

4.

5.

Preplant incorporated or preemergence applications
of alachlor at 3.36 or 4.48 kg/ha effectively
controlled yellow nutsedge.

With less than adequate rainfall (less than 2.5 cm
with in 7 to 10 days after treatment) preplant
incorporated alachlor treatments were more
effective than preemergence applications.

Alachlor did not effectively control yellow nutsedge
in high organic matter soils.

Reduction in yellow nutsedge shoot growth occurred
when alachlor was absorbed from the soil by the
tissue above the tuber.

Absorption of alachlor by roots did not reduce
plant growth.

Alachlor at 3.7 x 10"6 M inhibited growth of yellow

nutsedge shoots in Petri dish cultures.

65

10.

11.

12.

13.

14.

66
Alachlor at 3.7 x 10'”6 M killed newly emerging
nutsedge shoots in Petri dish cultures.
Alachlor at 3.7 x 10"6 M or 18.5 x 10'5 M did
not inhibit sprouting of yellow nutsedge tubers.
Depth of tubers in the soil did not affect
nutsedge response to alachlor.
Six weeks after treatment, alachlor that was still
present in a loam soil was primarily in the tOp
5 cm of soil.
Primarily acropetal and some basipetal Inc-
translocation occurred from 1“(t-alachlor
applications to small yellow nutsedge seedlings
with 2 to 4 cm shoots.
Alachlor was absorbed by lO-day-old plants and
translocated acrOpetally or basipetally to the
yellow nutsedge growing points killing the plants.
After lac-alachlor treatments on small nutsedge
seedlings, the 11+C-material moved from the shoot
and tuber rhizomes to the growing point, but the
majority of the 19c from 1L‘c-alaehlor applied to
the root did not move.
In tall (15 cm shoots) yellow nutsedge plants,

limited acropetal and no basipetal 1"‘C-translocation

occurred.

67
15. Postemergence alachlor applications do not control
yellow nutsedge, because alachlor is not trans-
located to the growing point.
16. Alachlor is metabolized rapidly in yellow nutsedge
to at least one water-soluble metabolite.
l7. Alachlor is an effective herbicide for controlling

yellow nutsedge in Michigan.

1.

2.

3.

7.

8.

9.

10.

11.

LIST OF REFERENCES

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Duncan, R. G., 0. N. Andrews, F. D. Timmsons, and
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74

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We
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soybeans. Weed Sci. 20:194-201.

75

88. Weed Society of America. 1970. Herbicide Handbook,
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APPENDICES

77

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78
Appendix B

Response of ellow nutsedge tuber sprouting
to 3.7 x 10' M alachlor treatments.

 

Exposure time to alachlor Percent sproutinga
(hr) (1 wk) (2 wk) (3 wk) (4 wk)

0 , 30.0 c 43.3 a 44.2 a 44.2 a

8 30.0 c 58.3 a 68.2 a 68.2 a

24 23.3 bc 38.7 a 38.7 a 38.7 a

48 16.7 abc 29.2 a 30.0 a 30.0 a

72 23.3 be 43.3 a 47.0 a 47.0 a

96 8.3 ab 30.5 a 33.3 a 37.5 a

120 6.7 a 22.0 a 31.2 a 36.2 a

144 30.8 c 47.0 a 48.7 a 48.7 a

 

aMeans within columns with common letters are not significantly
different at the 5% level by Duncan's Multiple Range Test.

 

79
Appendix C

New shoot emergence and growth of yellow nutsedge
from sand cultures c ntaining sprouted tubers
exposed to 3.7 x 10' M alachlor.

 

 

 

New shoot emergencea Shoot lengtha
Treatment 3 wk 4 wk 3 wk 4 wk
(No./culture) (No./culture) (cm/shoot) (cm/shoot)
No treatment 0.0 b 0.9 ab 11.9 b 19.1 a
Alachlor for 0.8 ab 2.1 a 2.0 c 10.0 b
1 week
Alachlor 0.0 b 0.0 b 0.3 o 0.9 o
continously

 

aMeans within columns with common letters are not significantly
different at the 5% level by Duncan's Multiple Range Test.

 

80
Appendix D

The separation by TLC of extracts from shootg
of ll-day-old yellow nutsedge 5 days after 1 CR
alachlor treatment.a

 

Rf
Treatment 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10.0
Water-soluble 0.0
fraction
Roxane-soluble 0.68
fraction
14C-alachlorb 0.68

 

3TLC plates were develOped in petroleum ether:chlcroform:95
percent ethanol (7:2:1; v/v/v).

bRing labeled 14c-alachlor was used as the standard.

81
Appendix E

The separation by TLC of water soluble alachlor
metabolites of ll-day-old yellow nutsedge 5 days

 

 

after C-alachlor treatment. a
, 77 RF
Treatment 0.0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1.0
water-soluble 0.33b

metabolites

 

8TLC plates were develOped in l-butanol .glacial acetic acid:
water (12: 3. 5)e

bRing labeled Inc-alachlor was used as the standard.and.had
an f of 0.60.

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