i

Wlllfillilllllllll‘illllllliIHIIHIIHIlHIHIIIIHHIWHI

mass 3 1293 00077 173

Y) 7‘. -‘
'L-err-I;;~ - ., ..

This is to certify that the
dissertation entitled
A CHARACTERIZATION OF THE CARROT ROOT PERIDERM

INCLUDING THE EFFECT OF HERBICIDES AND
LONG-TERM STORAGE

presented by

Lisa Ann O'Rear

has been accepted towards fulfillment
of the requirements for

Doctoral I degree in HQEII CU lture

<ZNQ77téé’KQ-J?QZ;Q£Z,
V

Major professor

Date (9C%c29/ /C/39/

MS U i: an Affirmative Action/Equal Opportunity Institution 012771

.3» $44.7 A-a'

 

MSU

 

 

 

RETURNING MATERIALS:
Place in book drop to

 

 

LIBRARIES remove this checkout from
“ your record. FINES W‘iII
be charged if book is
returned after the date
“fl" . stamped below.
.IaLi3:=if=-‘fi"6"“773
3 s20?
r Etyfso \

 

 

A CHARACTERIZATION OF THE CARROT ROOT PERIDERM
INCLUDING THE EFFECT OF HERBICIDES AND
LONG-TERM STORAGE

By

Lisa Ann O'Rear

A DISSERTATION

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

DOCTOR OF PHILOSOPHY

Department of Horticulture

1981

’f

G I ['2 0/32.»

ABSTRACT

A CHARACTERIZATION OF THE CARROT ROOT PERIDERM
INCLUDING THE EFFECT OF HERBICIDES AND
LONG-TERM STORAGE

by

Lisa Ann O'Rear

The periderm of the carrot root (Daucus carota L.) was

 

analyzed by chemical, physical and visual means to gain an under-
standing of its function and its development in various situations.
The studies were confined to those cell walls of the phellum which
resisted digestion with fungal pectinase. Microscopic examination
showed this membrane-like structure to be continuous over the outer
surface of the root. The weight per unit area of the isolatable
structure varied with cultivar from 470 to 790 ug/cm2 of which 7
to ll% was soluble in chloroform/methanol (CHCl3/Me0H). Thin layer
chromatography indicated that this lipid fraction contained 6 major
components, the most predominant of which remained at the origin and
co-chromatographed with fatty acids. Scanning electron microscopy
did not reveal any substantial wax surface structure.

The weight of the membrane per unit area decreased during
root growth of 'Scarlet Nantes' with reductions occurring in both
CHCl3/MeOH-soluble and insoluble portions, but no qualitative changes

in the wax composition were evident.

Lisa Ann O'Rear

A field trial of chlorpropham, ethofumesate, stoddard oil
and linuron sprays was performed but no significant effects on
'Danvers' were detected. Foliar applications of ethofumesate (2.24
kg/ha) to greenhouse-grown 'Scarlet Nantes' reduced both fractions
significantly and decreased the diffusion resistance of the root
tissue.

Total periderm membrane weight of roots stored at 0 C
increased over a 6-month period. Soluble lipid per unit area
increased in ’Spartan Bonus' but not in 'Tito' during this time.
The insoluble fraction increased dramatically in both cultivars.
Quantitation of suberin in the insoluble material by an assay of
the hydrogenolysis products, C16 and C18zl diol showed suberin syn-
thesis to occur between 0 and 3 months.

The artificial removal of periderm components showed the
suberized walls to be a barrier to water loss from the root, but
were substantially reduced in effectiveness when the soluble lipids

were extracted.

DEDICATION

to Rick

ii

 

ACKNOWLEDGEMENTS

I wish to thank Dr. James A. Flore for serving as my major
professor and for providing guidance throughout my degree program.
Drs. J. N. Cash, R. C. Herner, A. R. Putnam and M. N. Zabik partici-
pated on my advisory committee and provided valuable assistance.

The following persons also contributed to my endeavors:

Dr. B. B. Dean, Dr. S. Flegler, Dr. L. R. Baker, Bill Chase and
Viv Shull.

My fellow graduate students made it bearable.

iii

TABLE OF CONTENTS

List of Tables .

List of Figures.

LITERATURE REVIEW .

SECTION I A CHEMICAL AND DEVELOPMENTAL CHARACTERIZATION
OF THE CARROT ROOT PERIDERM. .

Abstract

Introduction . . .

Materials and Methods .
General.
Plant material and culture
Isolation of periderm membrane .
Collection of soluble lipids.
Thin layer chromatography.
Preparative layer chromatography
Light microscopy. .
Scanning electron microscopy.

Results and Discussion.
Light microscopy.
Isolation of membrane and extraction of
soluble lipid. . . .
Scanning electron microscopy. .
Thin layer chromatography (TLC).
Developmental studies . .

Summary and Conclusions

Literature Cited.

SECTION II ALTERATION OF THE CARROT ROOT PERIDERM BY ETHO-
FUMESATE . .

Abstract

Introduction . . .

Materials and Methods .
General.
Plant material and culture .
Lipid extraction and quantitation .

iv

Page
vi

vii

Thin layer chromatography. .
Diffusion resistance measurements .
Results and Discussion. .
Field trial . .
Greenhouse trial of ethofumesate
Qualitative analysis
Diffusion resistance
Summary and Conclusions
Literature Cited.

SECTION III THE FUNCTION AND CHEMISTRY OF THE CARROT ROOT
PERIDERM IN LONG- TERM STORAGE . .

Abstract

Introduction . . .

Materials and Methods .
General. .
Desiccation studies.
Storage experiments.

Results and Discussion.
Desiccation experiments
Storage studies .

Summary and Conclusions

Literature Cited.

APPENDICES
A Desiccation of Five Carrot Lines . .
B The Effect of Herbicides on Red Beet and Sugar
Beet. . . .
C Modified Half- Strength Hoagland' 5 Solution .
D Herbicide Data . . .
BIBLIOGRAPHY.

65
69
75

77

LIST OF TABLES

Table Page
1 Comparison of periderm components from carrot lines. l8

2 Quantitative fractionation of soluble lipid from MSU
1410 B by thin layer chromatography in benzene . 22

3 Effect of herbicides on the concentration of soluble
periderm lipids in 'Danvers' . . . . . . . 36

4 Comparison of periderm extraction techniques . . . 38

5 The effect of ethofumesate on the periderm composition
of 'Scarlet Nantes' . . . . . . . . . . 39

6 Diffusion resistance of carrot roots lacking periderm
or soluble periderm lipids . . . . . . . . 54

7 Periderm composition of 'Tito' and '5. Bonus' in long-
term cold storage . . . . . . . . . . . 55

8 Suberin diol content of '5. Bonus' periderm in long-
term storage. . . . . . . . . . . . . 59

9 Desiccation of 5 carrot lines at D C. . . . . . 66

lo Desiccation of 5 carrot lines at 5 C. . . . . . 67

ll The effect of herbicides on periderm membrane weight
and desiccation in red beet. . . . . . . . 7D
12 Desiccation rates of herbicide-treated sugar beets . 7l

13 Modified, half-strength Hoagland's Solution . . . 73

l4 Herbicide nomenclature, formulation and manufacturer 75

vi

LIST OF FIGURES

Cryostat section (7p) of 'A Bak' root stained with
Sudan III and IV, l25 x. . . . . .

Sudan-stained, cryostat sections of periderm isolated
with pectinase. A 7p section of material treated
for 48 hr., 125 x B. 10p section following 96 hr.
treatment, 250 x . . . . . . . .

Scanning electron micrograph of 'Tito' root surface,
860 x. A. Normal surface. 8. Surface after wiping
with CHCl3/Me0H-soaked cotton.

Thin layer chromatogram of soluble lipids from peri-
derm membrane of 5 carrot lines and leaf wax from
cabbage. Developing solvent = benzene.

Changes in periderm and root fresh weight over time
in greenhouse-grown 'Scarlet Nantes'. Each point
is the average of 4 replicates of lD-l5 roots.
Vertical bars indicate standard errors.

Changes in the composition of the periderm membrane
during the growth of 'Scarlet Nantes'. Each point
is the average of 4 replicates of 10-15 roots.
Vertical bars indicate standard errors.

Thin layer chromatogram of soluble lipids from
isolated 'Scarlet Nantes' periderm at various
harvest dates. Number of days after sowing is
indicated below each sample. Developing solvent
= benzene . . . . . . . . . .

Thin layer chromatogram of soluble periderm lipids
from 250 cm2 surface of 'Scarlet Nantes'. Develop-
ing solvent = benzene. A. Control B. Ethofume-
sate-treated. (2.24 kg/ha) C. Cabbage leaf wax

Desiccation patterns of carrot roots with and without
periderm or soluble lipids. A. Storage in l00%
RH. B. Storage in 88% RH. Temperature =
Vertical bars indicate standard errors.

vii

Page

15

16

20

21

24

25

26

41

52

Figure Page

l0 Thin layer chromatogram of soluble periderm lipids
from '5. Bonus' following various storage periods
A. Cabbage leaf wax. B. 0 mos. C. 3 mos.
D. 6 mos. Developing solvent = benzene. . . . 56

ll Gas chromatogram of THC ethers of suberin diols from

'5. Bonus'. Column = 3% SE-30. N2 flow rate =
25 ml/min. Temperature = 220 C. . . . . . . 58

viii

LITERATURE REVIEW

The peripheral tissues of higher plants are separated from
the external environment by membrane-like coverings. Most aerial
organs are sheathed by the cuticle, whose structural component is
the lipid polymer, cutin. Associated with the cutin matrix are
embedded and surface (epicuticular) waxes. Cellulose and pectin
occur in the area nearest the cell wall (34). The surface cells of
underground organs (25), wound-healed parts (7, 24) and bark tissue
(l0) are served by a distinct but similar polymeric material, suberin.
In the suberized tissues such as the periderm of roots, the suberin
exists as a complex associated with the cell wall and is not deposited
on the surface as is the cuticle. In addition, suberized cells have
been located in various internal tissues (8, 30, 35, 39).

Many functions of cuticles and suberized plant tissues are
known and others have been postulated. The cuticle is able to minimize
transpirational water loss from leaves and fruit due to the wax com-
ponent (l4, l5, 36, 40). It is often noted that epicuticular waxes
in particular are responsible for this phenomenon (36). The resis-
tance of potato wound periderm to water loss was also attributed to
the waxes embedded within the suberin complex (42). The presence of
suberized cells in internal tissues implies the movement of substances
within the plant may be regulated. For example, the endodermis of
roots acts as a barrier to apoplastic movement of ions across the

l

the radius of the root (4). The role of suberized gland cells (30)

and bundle sheath cells (27) remains obscure. The wax component of the
cuticle inhibits the penetration of chemicals (2, 15, 32, 40) more

so than does the thickness of the cutin matrix (31, 40). In parti-
cular epicuticular wax of leaf cuticle has been shown to influence

the wettability of leaves and thus the penetration of ions (2, 32,

29). A qualitative analysis of surface wax detected differences

among chemical groups in their ability to impede water movement.
Alkanes and aldehydes were the most effective compounds in these
studies (2, 14).

Cuticles and suberin-enriched tissue may be isolated for
study by any one or a combination of several methods including treat-
ment with ammonium oxalate, oxalic acid, ZnCl2 or fungal pectinase
and cellulase. Suberin, however, is difficult to obtain in pure form
due to its close association with the cell wall.

Following isolation of the membranes as described above,
additional extractions with carbonate solutions and organic solvents
can be used to further purify the cutin or suberin. The monomeric
components of the polymers are obtained by alkaline hydrolysis,
transesterification or hydrogenolysis, each reaction yielding dif-
ferent derivatives (i.e., varying degrees of reduction) of the
original components (19). Cutin is found to be largely constructed
of 16— and l8-carbon carboxylic acids with their mono-, di- and
trihydroxy and epoxy constituents. The predominant component is
dihydroxy palmitate (18). Small amounts of phenolics are present

(38), but the polymer is 70-90% aliphatic. Suberin, on the other

hand, characteristically contains a higher proportion of phenolics,
and its fatty monomers are dominated by w-hydroxy oleic, w-hydroxy
palmitate, their dicarboxylic analogs and alcohols of 20 or more
carbons (3, ll, 21, 25). Like the cuticle, suberized periderm tissue
contains wax-like compounds (12, 33).

Detailed characterizations of periderm suberin from several
underground plant organs are available (25). In the case of carrot,
the suberin residue released by hydrogenolysis of the periderm was
found to be 26% phenolic and 9% aliphatic by weight. 0f the ali-
phatic constituents, dicarboxylic acids accounted for 24% and the
major homologs in decreasing abundance were c18zl’ C16’ and C18.
w-hydroxy acids (predominantely C18:1’ C16, C18, and C20) comprised
31% of the aliphatic fraction. Fatty acids and fatty alcohols were
14 and 3% of the total respectively; the major monomer in both cases
was C22.

- Soluble lipids (waxes) were present in concentrations of 2
to 32 ug/cm2 in one study of the periderm 0f subterranean organs (12).
The suberin-associated soluble lipids of carrot are 50% fatty acid,
34% primary alcohol with 5% accounted for by alkanes and wax esters.
The major homologs of acids and alcohols were C18 to C24 while those
of the alkanes C25 to C31.

A proposed structure for the suberin complex has a phenolic
matrix covalently attached to the cell wall with aliphatic compounds
bonded to the phenols (20). The polymer is deposited in the primary

cell wall near the cytoplasmic side. A lamellar structure of light

and dark bands as viewed with the electron microscope suggests that
waxes alternate with the suberin polymer (8, 42).

The synthesis of suberin has been studied in wounded tissues
(6, 7). Potato tuber pieces incorporated 14C-labelled acetate and
oleic acid into a chloroform-insoluble residue, subsequently found
to contain labelled urhydroxy acids and dicarboxylic acids. The
synthesis of these compounds reflected the rate of increase in dif-
fusion resistance, a measure of suberin deposition. The induction
of suberin synthesis in wound situations required 2 to 3 days and
apparently involved a trigger of unknown identity (6). Tissues which
normally were covered by a cuticle (and thus, cutin) were found to
synthesize suberin upon wounding (7). It is assumed that the syn-
thetic pathways are identical for wound and natural suberin since
their chemical composition is the same.

The effect of certain agrichemicals on cuticular lipids has
been a concern of plant scientists since the publication of Dewey
et al in 1956 (9). Their report implicated trichloroacetic acid as
an inhibitor of epicuticular wax deposition in pea. Since then,
several other compounds have been found to have similar effects on
various species. Most attention has been focused on the thiocarba-
mate herbicides, such as EPTC and diallate. The consequences of
reduced leaf wax following applications of these and other chemicals
include increased uptake of foliar applied compounds (5) and increased
cuticular transpiration (28). The resultant increased wettability
of the leaves is likely due to the disruption and/0r reduction in

epicuticular wax fine structure (16, 28). The chemical composition

of the wax may also be altered. Typically alkanes, secondary alcohols
and ketones are decreased (13, 22, 28, 45) while esters increase

(13, 28). These qualitative changes in wax composition of treated
plants point to an inhibition of the elongation/decarboxylation of
fatty acids by EPTC (22) and diallate (43) as the probable mode of
action. The consequences of decreased alkanes may include enhanced
penetration of the cuticle as discussed previously.

In at least one report (45), cuticle thickness has been
affected by EPTC-inhibition of the fatty acid-to-alkane sequence of
reactions of wax synthesis. Flore (13) found no change in cutin
monomers following EPTC treatments to cabbage, even though wax syn-
thesis was severely inhibited. Soliday et a1 (42) have examined the
ultrastructural aberrations of wound periderm caused by trichloro-
acetic acid. The lamellar arrangement of wax deposits was disrupted
by the treatment and total soluble lipids were decreased. The prac-
tical aspect of these alterations in structure and chemistry of
potato wound periderm was shown to be a decrease in diffusive resis-
tance of the tissue. This last report is of particular relevance
for stored root crops which can suffer serious losses from desicca-
tion. Any decrease in the protection afforded by the periderm will
probably further compound the problem. Chlorpropham, one of the
herbicides recommended for carrot in the state of Michigan has been
noted as an inhibitor of wax deposition (22) but not of suberization
in wounded potato tubers (37). No information is available to show
if the carrot periderm is being adversely affected by the use of

this carbamate herbicide or other chemicals.

In addition to chemicals, cuticular waxes are sensitive to
their external environment. Generally, light increases the quantity
of foliar wax while higher temperatures and higher relative humidi-
ties tend to decrease deposition (1.44). In general, the responses
are advantageous either for allowing greater cuticular transpiration
or for greater protection, whatever the environmental situation
dictates. The interaction of all factors will determine not only
the quantity of cuticular lipids, but also their chemistry (33) and
morphology (l, 44). As noted previously, these factors must be
considered in the role of wax as a barrier to penetration (2, 14).
The environment of the root is also variable with respect to moisture
and temperature. Periderm lipids of roots may be expected to respond
to physical extremes particularly if their biosynthetic pathways
are identical or similar to those of leaf waxes.

The periderm of economically important root crops, such as
the carrot is an interesting topic of study. Carrot roots are in
secondary growth before the first true leaves emerge and therefore
the periderm is a factor in the function of the root as an organ of
uptake. The suberized cells in particular are likely to affect the
movement of substances into and out of the root, both during the
development of the plant in the field and later during post harvest
handling and storage.

Although information on carrot periderm chemistry is beginning
to accumulate, the literature is for the most part descriptive in
nature. This dissertation in three sections reports a further char-

acterization of the carrot root periderm and follows its development

 

from a small plant through to maturity and then into long-term cold
storage. The effect of selected herbicides was included to evaluate

their effect on periderm development.

SECTION I
A CHEMICAL AND DEVELOPMENTAL CHARACTERIZATION
OF THE CARROT ROOT PERIDERM
Abstract

Periderm from the roots of carrot (Daucus carota L.) was

 

isolated enzymatically for chemical and structural examination. The
outer tranverse walls of the phellum layer formed a continuous,
external membrane containing chloroform/methanol-soluble lipids.
Separated by thin layer chromatography, these lipids contained at
least six'major chemical groups, the most abundant of which co-chroma-
tographed with fatty acids. Scanning electron microscopy revealed

the absence of surface fine structure. The periderm membrane
decreased in weight with development of the root, attributed to
reductions in both chloroform-methanol soluble and insoluble material
per unit area. The results are discussed in the light of current

knowledge of the plant cuticle structure and function.

Introduction

 

The roots of crops such as carrot, beet and turnip are covered
by periderm tissue at maturity (5). The lipid polymer, suberin, is
a characteristic component of the periderm cell walls (13) and has
been partially characterized in 5 root crops (14). In addition, wax-

like materials have been collected and identified from these plants

(7). Suberin-containing portions of the periderm can be separated
from adjacent tissue with enzymatic methods commonly employed to
isolate cuticles from aerial plant parts (14).

The cuticular membrane is formed in part by a lipid polymer,
cutin (17). Cellulose and pectin lie between a layer of cutin and
the cell wall and thus the cuticle can be released by digestion with
cellulase and pectinase. The cutin matrix is impregnated with wax
which may or may not be present as elaborate fine structure on the
outer surface (epicuticular wax). The chemistry of cutin and suberin
are similar in that both are composed in part of long chain fatty
acids, hydroxy fatty acids and related aliphatic compounds (13).
Phenolic constituents are present in both polymers (23) but to a
greater extent in suberin (11). Many of the soluble lipids associated
with suberin are those typical of cuticular waxes, i.e., alkanes,
esters, alcohols and fatty acids (7, 26).

The structure, location and chemistry of the cuticle limits
plant transpirational water loss and provides a barrier to the uptake
of foliar-applied chemicals. In addition, the cuticle serves as a
protective barrier to insect and disease infestation (17). The
suberized cell walls of the carrot periderm may also function as the
primary barrier to substances moving into and out of the root.
Although the suberin and its associated soluble lipids exist as a
lamellar structure within the cell wall, and are not extracellular
as in cutin, the periderm probably plays a similar role in preventing

water loss from internal tissue (4, 25). Like the cuticle (3, 10,

10

24) it is the waxes of the suberin complex which appear to constitute
the greatest impedence to water vapor diffusion (25).

An anatomical description of carrot root development was
published by Esau (6). The activity of the cambium and secondary
tissues displaces the cortex early in the life cycle of the plant
and the periderm forms the outer. Although the chemical composition
of carrot suberin and its associated waxes has been partially ana-
lyzed for mature roots, there is no developmental study of the peri-
derm with respect to its chemistry. The following investigations
were initiated to form a basis for physiological studies on the
carrot periderm, particularly the portion forming the outermost sur-
face 0f the root. Also reported are findings on changes in qualita—
tive composition of these suberin-enriched cell walls during growth

in a greenhouse situation.

Materials and Methods
General.--Fungal pectinase was obtained from United States
Biochemical Corporation, Cleveland, Ohio and the Tissue Tek II O.C.T.
Compound from Lab-Tek Products, Naperville, Illinois. Filter paper
and thin layer plates were pre-washed in the solvent or solvents to

be used with them. All solvents were glass-distilled.

Plant material and culture.--Carrots for the characterization

 

work were grown at the Michigan State University Muck Soils Research
Farm in Laingsburg, Michigan. 'Tito', 'A Bak', MSU 1414 B, MSU 1410
B and MSU 1389 B were obtained from the Department of Horticulture's

carrot breeding program. Seeds of 'Gold Pak', 'Danvers' and 'Spartan

11

Fancy' were purchased from Joseph Harris C0., Rochester, New York.
All were grown with standard cultural procedures.

Seeds of 'Scarlet Nantes' were obtained from Harris for the
development study. These were sown in a greenhouse in lO-liter
plastic pots filled with vermiculite. This medium greatly facilita—
ted the harvest and subsequent analyses as the particles were easily
rinsed from the root with minimum damage to the periderm. Fertiliza-
tion was provided by weekly applications of half-strength modified
Hoagland's (Appendix C) and Peter's 20-20-20 (rate = 200 ppm N).

No other chemicals were applied. A randomized complete block design
with 4 blocks was utilized. Twenty to 30 roots formed an experimental
unit. Six equally distributed sampling dates began 78 days after
sowing and ended on day 147. At each harvest, average root fresh
weight and total periderm membrane weight per unit area were deter-
mined. The periderm membrane was further divided into its chloroform-
soluble and insoluble components which were also quantitated on a

unit area basis. The soluble lipids were examined by thin layer

chromatography (TLC).

Isolation of periderm membrane.--Peels from mature carrot

 

roots were boiled for 15 minutes in distilled water and drained.
These were treated with a 1% (w/v) solution of fungal pectinase in
citrate-phosphate buffer (pH 3.8). Complete removal of cellular
debris (as determined by microscopic examination) usually occurred
after 4 days with mechanical agitation and required at least 1

change of solution during this period. For accurate determination

12

of membrane weight per unit area, discs of 6-mm diameter were punched
with a cork borer and excised from the root surface with a razor
blade in preparation for enzyme treatment. Following digestion of
the excess internal tissue, the resultant cell-free membranes were

thoroughly rinsed, drained and air-dried.

Collection of soluble lipids.--Isolated, air-dried membranes
were refluxed for 2 hours in chloroform/methanol (CHC13/MeOH 2:1
v/v). The solvent containing dissolved lipids (waxes) was filtered
through Whatman no. 1 paper. This procedure was repeated with clean
solvent and the extracts combined. Extracts were concentrated to
approximately 3 m1 under a stream of N2, transferred to screw-capped
test tubes for complete drying and sealed under N2 until used in
qualitative analyses. The membranes were weighed before and after

extraction for calculation of soluble lipid weight.

Thin layer chromotography.--The extracts of the five lines

 

in the characterization study were diluted to 10 mg/ml. Those of

the development work were adjusted to reflect 250 cm2 area extracted/
ml based on their calculated weight/cmz. The lipids were applied as
spots (100 pl) or 0.5 mm-long bands (200 pl) to 0.25 mm silica gel

G plates, prepared as described by Flore and Bukovac (8). Cabbage
leaf wax of known composition (2, 8) was applied with the lipids of
the 5 lines for comparison. Development in benzene was followed
with a 50% sulfuric acid spray. Plates were immediately heated at
100 C for approximately 5 minutes and checked for color reactions

before allowing to char.

13

Preparative layer chr0matograpby.--F0r quantitative analysis
of major wax fractions by weight, larger (10 mg) quantities of soluble
lipid from MSU 1410 B were applied as a l6-cm band to 0.5 mm silica
gel G. Plates were again developed in benzene then exposed to iodine
vapors for detection of the separated fractions. Prominant bands
were outlined to facilitate location after the stain evaporated.

Each band was scraped from the plate and extracted with 10 ml CHCl3/
MeOH (2:1 v/v) at 60 C for brief periods. The slurry was washed
through Whatman no. 1 paper into clean tubes. The chromatography of
each fraction was repeated and the bands collected as before but
into tared tubes. Solvent was evaporated under N2 and the tubes

weighed.

Light microscopy.--Isolated membrane discs and fresh carrot

 

root pieces (2 or 3 mm) containing periderm were surrounded with
Tissue-Tek II and quick frozen on metal stubs in an International
Harris Model CTD cryostat. Following the method of Norris and Bukovac
(19), microtome sections 6—12 u thick were cut and mounted on glass
slides at -20 C. Slides were flooded with a saturated solution of
ethanolic Sudan III and IV for 30 seconds. After rising and air-
drying, cover slips were affixed with 1% (v/v) phenol in 1:1 (v/v)

water and glycerol.

Scanning electron microscopy.--Small (2 to 3 mm) pieces of
tissue were cut from the surface of 'Tito' roots, frozen on dry ice

and lyophilized. Some sections received a pre-treatment which

l4

consisted of wiping the surface with CHCl3/Me0H-soaked cotton prior
to freezing to remove any superficial wax deposits. For microscopy,
sections were coated with approximately 20 nm gold. Photographs were

taken off of a JEOL Model JSM 35C instrument operated at 15 kV.

Results and Discussion

Light microscopy.--A section from the mid section of mature

 

carrot root surface ('A Bak') is pictured in Figure l. Rectangular-
shaped phellum cells comprise the outermost layer of periderm. The
darkened walls of these cells stained positively with Sudan, a stain
that has been used in histological studies to detect cutin and suberin
(19, 22). Carrot periderm is known to contain suberin (l4) and

Figure 1 illustrates the location of suberized cells. Note that in
some areas the stained cells are two-deep, but the heaviest staining
area is the outermost surface.

Periderm cells containing suberin are expected to resist
digestion by pecteolytic enzymes and this indeed was the case with
carrot root. Figure 2 illustrates the condition of the phellum layer
after 2 (2a) and 4 (2b) days of enzyme treatment. The entire outer
row persists after the first 48 hours but the anticlinal and inner
transverse walls are further degraded with continued treatment. The
result is that a continuous membrane is formed mainly by the outer
transverse wall of the phellum. This wall is isolated intact from
the underlying tissue.

This observation leads to the first parallel (with some

restrictions) that can be drawn between the carrot periderm and a

 

Figure l.--Cryostat section (7p) of 'A Bak' root stained with Sudan
III and IV, 125x.

 

Figure 2.--Sudan-stained, cryostat sections of periderm isolated with
pectinase. A. 7 u section of material treated for 48 hr,125x.
B. 10 u section following 96 hr of treatment, 250x.

l7

characteristic leaf cuticle. Both the carrot root and the typical
leaf are covered by an outer, continuous membrane structure, however
in the case of the leaf, the cuticle is formed on top of the epi-
dermal cell walls. The cutin-containing matrix is cleanly separated
from these walls upon pectinase treatment (10, 19, 20). Adjacent
phellum cells of the carrot periderm release their outer walls
intact upon digestion of the tissue. The suberin contained within
these walls probably forms continuous bands across the areas of the
radial cell walls (4.25) and thus lends continuity to the surface

of the root.

Isolation of membrane and extraction of soluble lipid.--The
weight per unit area of the isolated membrane was found to vary with
cultivar and range from 470 to 790 pg/cm2 (Table 1). Although some
of the observed differences could be due to environmental and loca-
tion effects, significant variation was shown among 5 lines grown in
the same experimental plot. The proportion of the total membrane
which was extracted in CHCl3/MeOH is listed in Table l as '% soluble
lipid'. The hot solvent mixture and the reflux extraction procedure
utilized was designed to remove all lipids and other lipophillic
organic compounds other than those covalently bound as part of the
suberin polymer (14). The range of soluble lipid concentrations is
not as wide as that of total membrane weights, but remains rather

constant among cultivars, taking on a value of 7 to 11%.

Scanningelectron microscopy.--Scanning electron microscopy

was employed to determine if soluble lipid fine structure typical of

18

TABLE 1.--Comparison of periderm components from carrot lines.

 

 

 

Total
Cultivar membrane wt. % soluble
(us/cmZ)

'Spartan Fancy' 470 i 50 7.0 i 2
'Danvers' 640 i 60 10.0 i 3
'Gold Pak' 79- i 70 8.0 i l
MSU 1389 32 710.1y 8.3a
'A Bak' 710a 9.6a
MSU 1410 B 640b 11.0a
'Tito' 630b 10.6a
MSU 1414 8 600b 10.7a

 

2This line and those which follow were grown in the same experimental

plot. Values represent the average of 3 reps.

yMean separation within columns by Duncan's Multiple Range Test, 5%

level.

19

leaf and fruit waxes (2, l6) existed on the outside of the root.
Micrographs (Figure 3) revealed no regular pattern of wax crystals.
The surface debris and ridge-like matter was not greatly reduced by
CHCl3/MeOH extraction. Typical leaf and fruit wax compounds would
have been removed by this treatment (1, 19, 21). The superficial
structure seen in Figure 3 is evidently not a lipid or at least not
a free (soluble) lipid form. These observations illustrate a major

difference between roots and cuticularized organs.

Thin layer chromatography (TLC).--In order to verify the
existence of typical leaf wax compounds, TLC was performed on the
soluble lipids from isolated membrane material of MSU 1410 B. A
non-polar solvent, benzene, was selected to separate out hydrophobic
classes from the crude extract (2, 8). In Figure 4, cabbage wax
previously characterized is used for comparison with the carrot
extract. Non-polar compounds, migrating above the origin are shown
to constitute a portion of the soluble lipids, in the periderm mem-
brane and the six most prominant spots were collected from prepara-
tive layer plates for gravimetric analysis. The most polar fraction,
6 is also the predominant one (Table 2), confirming the findings of
Espelie et a1 (7). Relatively small amounts of very non-polar classes
(i.e., fractions 1 and 2) were recovered. The opposite was found to
be true of cabbage wax in at least one study (8) where alkanes made
up 26% of the and fatty acids only 3.5%. Collectively then, the
carrot periderm lipids are relatively more polar than the cabbage

wax and would be expected to lend different permeability

20

1
75
l
1

 

Figure 3.--Scanning electron micrograph of 'Tito’ root surface, 860x.

A. Normal surface. B. Surface after wiping with CHCl3/MeOH-
soaked cotton.

alkane

ester
ketone

aldehyde

2° alcohol
ketol

10 alcohol

 

fatty acid

Figure 4.--Thin layer chromatogram of soluble lipids from periderm
membrane of 5 carrot lines, and leaf wax from cabbage.
Development in benzene.

22

TABLE 2.--Quantitative fractionation of soluble lipid from MSU 1410 B
by thin layer chromatography in benzene.

 

 

Fractionz % of
number Rf total
1 1.0 1.52

2 0.95 4.57

3 0.61 2.03

4 0.33 4.06
0.10 14.21

6 0.00 65.49

 

zFrom Figure 4.

23

characteristics to the suberized cell walls. The contribution of
various chemical classes to the effectiveness of leaf wax as a barrier
to penetration has been documented (3, 9) but similar studies have

not been performed for suberin-associated waxes.

Developmental studies.--Data taken during carrot growth

 

enabled us to correlate periderm components with root development.
Total periderm membrane weight/cm2 and root fresh weight are plotted
against time in Figure 5. The weight of the isolated membrane is
seen to decrease linearly with time as the size of the root increases.
Figure 6 illustrates the change in the soluble lipids and insoluble
, portion of the developing root periderm. The weight/unit area of
both components decreased during the course of the study, the soluble
portion at a slightly greater rate. The crude (unextracted) membrane
was 6.6% soluble at day 78 and 5.8% soluble at termination. Since
the root is rapidly expanding over the period in which these observa-
tions were made, it is likely that the synthesis of periderm wall
components did not keep pace with the increase in the surface area of
the root. The data in this study supports this explanation since
all membrane components decreased in quantity and at approximately
the same rate.

Cuticular lipids have been shown to undergo changes in com-
position during maturation (4, 16), however, qualitative differences
in wax between sampling dates were not observed with TLC (Figure 7).

It should be pointed out however, that this technique is not sensitive

24

0
J.

r t

1.00 ‘1- ’ 1b 50

H ROOT FRESH WT.
m MEMBRANE WY.

‘0‘ 0.80" db‘O

E

i?

1;: o.eo~- "3°

5

O ' .

g 0'40" " 1r20
0.20-1- 010

 

 

 

oaL—t: : t

: O
O 75 100 125 150
DAYS AFTER SOWING

Figure 5.--Changes in periderm and root fresh weight with time in
greenhouse-grown 'Scarlet Nantes'. Each point is the
average of 4 replicates of 10-15 roots. Vertical bars
indicate standard errors.

(6) 'lM 14838:! 1.008

25

 

 

 

 

ooh—n 4 : 4. 000
so» «700
m sown:
00—0 INSOEUILE
A ‘0 do 6002
0)
"s o
\ I"
5 1:":
a)
a so «500 g
A 31
M
E!
g ..
r.
3 20“ «400 a
“P '2
O
i»
‘0 0 "30°
0 <L—g‘ : t 4 O
o ’ 75 100 125 150

DAYS AFTER SOWNG

Figure 6.--Changes in the composition of the periderm membrane
during the growth of 'Scarlet Nantes'. Each point is
the average of 4 replicates of 10-15 roots. Vertical
bars indicate standard errors.

26

0.40

0.26
0.20

0.10

0.00

 

78 92 105 119 133 147

-Figure 7.—-Thin layer chromatogram of soluble lipids from isolated
'Scarlet Nantes' periderm at various harvest dates. Number
of days after sowing is indicated below each sample. Develo-
ping solvent = benzene. Each band contains the soluble
lipids from 250 cm2 of root surface.

27

enough to detect subtle changes in concentration of the different

chemical groups from one date to another.

Summary and Conclusions

The outer cell wall of the carrot root periderm forms a
continuous membrane over the root surface. Soluble lipids comprise
approximately 10% of the membrane and the bulk of them are relatively
polar. These components appear to be entirely embedded rather than
superficial. However, synthesis of the suberized portions of the
periderm and the associated soluble lipids does not proceed at the
same rate as root expansion as seen by a progressive decrease in the
weight per unit area of the membrane.

Differences in membrane weight between cultivars suggest the
possibility of genetic manipulation. As with cuticular waxes, chemi-
cal and environmental manipulations of the waxes may be possible.

The physical and chemical properties discussed here suggest
a protective role for this portion of the root periderm. When com-
pared to a leaf cuticle, however, the periderm membrane lacks some
features which are very important to the function of the leaf cuticle,
i.e., surface wax fine structure and a high concentration of very
'hydrophobic lipid classes. The permeability characteristics of the
carrot periderm and the contribution of individual components requires
additional study before further analogies can be drawn between the

protective role of the cuticle and the suberized tissues.

10.

11.

28

Literature Cited

 

Bain, J. M. and D. McG. McBean. 1967. The structure of the
cuticular wax of prune plums and its influence as a water
barrier. Aust. J. Biol. Sci. 20: 895-900.

 

Baker, E. A. 1977. Sources of variability in plant epicuticular
waxes and some effects on foliar penetration. Ph.D. thesis,
Univ. of Bristol.

Baker, E. A. and M. J. Bukovac. 1971. Characterization of the
components of plant cuticles in relation to the penetration
of 2.4-0. Ann. Appl. Biol. 67: 243-253.

 

Dean, B. B., P. E. Kolattukudy and R. W. Davis. 1977. Chemical
composition and ultrastructure of suberin from hollow heart
tissue of potato tubers (Solanum tuberosum). Plant Physiol.
51: 1008-1010.

Esau, K. 1977. Anatomy of Seed Plants, 2nd ed. John Wiley
and Sons, Inc., New York.

Esau K. 1940. Developmental anatomy of the fleshy storage
organ of Daucus carota L. Hilgardia 13: 175-226.

Espelie, K. E., N. Z. Sadek and P. E. Kolattukudy. 1980.
Composition of suberin-associated waxes from the subterranean
storage organs of seven plants. Planta 148: 468-476.

Flore, J. A. and M. J. Bukovac. 1978. Pesticide effects on
the plant cuticle: III. EPTC effects on the qualitative
composition of Brassica oleracea L. leaf cuticle. J. Amer.
Soc. Hort. Sci. 103: 297-301.

Grncarevic, M. and F. Radler. 1967. The effect of wax compo-
nents on cuticular transpiration-model experiments. Planta
75: 23-27.

Haas, K. and J. Schbnherr. 1979. Composition of soluble cuti-
cular lipids and water permeability of cuticular membranes
from Citrus leaves. Planta 146: 399-403.

Kolattukudy, P. E. 1980. Biopolyester membranes of plants:
cutin and suberin. Science 208: 990-1000.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

29

Kolattukudy, P. E. 1978. Chemistry and biochemistry of the
aliphatic components of suberin, p. 43-84. In G. Kahl, (ed.)
Biochemistry of wounded plant tissues. de Gruyter and Co.,
New York.

Kolattukudy, P. E. 1977. Lipid polymers and associated phenols.
p. 185-257. In F. Loewus and V. C. Ruenckles (eds.) Recent
advances in Phytochemistry, V01. 11. Plenum Press, New York.

Kolattukudy, P. E., K. Kronman and A. J. Poulose. 1975. Deter-
mination of the structure and composition of suberin from the
roots of carrot, parsnip, rutabaga, turnip, red beet and
sweet potato by combined gas-liquid chromatography and mass
spectrometry. Plant Physiol. 55: 567-573.

 

Leece, D. R. and A. L. Kenworthy. 1972. Influence of epicuti-
cular waxes on foliar absorption of nitrate ions by apricot
leaf disks. Aust. J. Biol. Sci. 25: 641-643.

Long, S. M. 1980. Cuticle development and incidence of russet
on 'Golden Delicious' apple as influenced by subclone suscep-
tibility and shelters. M.S. thesis, Michigan State University,
East Lansing.

Martin, J. T. and B. E. Juniper. 1970. The Cuticles of Plants.
St. Martin's, New York.

Norris, R. F. and M. J. Bukovac. 1972. Influence of cuticular
waxes and penetration of pear leaf cuticle by l-NAA. Pesti.
Sci. 3: 705-708.

Norris, R. F. and M. J. Bukovac. 1968. Structure of the pear
leaf cuticle with special reference to cuticular penetration.
Amer. J. Bot. 55: 975-983.

 

Orgell, N. H. 1955. The isolation of plant cuticle with pectic
enzymes. Plant Physiol. 30: 78-80.

Possingham, J. V., T. C. Chambers, F. Radler and M. Grncarevic.
1967. Cuticular transpiration and was structure and composi-
tion of leaves and fruit of Vitis vinifera. Aust. J. Biol.
Sci. 20: 1149-1153.

Priestly, J. H. 1921. Suberin and cutin. New Phytol. 20:
17-29.

Riley, R. G. and P. E. Kolattukudy. 1975. Evidence for
covalently attached coumaric and ferulic acids in cutins and
suberins. Plant Physiol. 56: 650-654.

24.

25.

26.

30

Schbnherr, J. 1976. Water permeability of isolated cuticular
membranes: The effect of cuticular waxes on diffusion of
water. Planta 131: 159-164.

Soliday, C. L., P. E. Kolattukudy and R. W. Davis. 1979. Chemi-
cal and ultrastructural evidence that waxes constitute the
major diffusion barrier to water vapor potato tuber (Solanum
tuberosum L.) Planta 166: 207-214.

Tulloch, A. P. 1976. Chemistry of waxes of higher plants. p.
235-287. In P. E. Kolattukudy (ed.) Chemistry and biochemis-
try of natural waxes. Elsevier, Amsterdam, NY.

SECTION II

ALTERATION OF THE CARROT ROOT PERIDERM BY
ETHOFUMSATE
Abstract

Soluble periderm lipids of carrot (Daucus carota L. var

 

'Danvers') were not quantitatively affected by foliar sprays of
linuron (3-(3, 4-dichlorophenyl)-l-methoxy-l-methylurea, 1.12 kg/ha),
chlorpropham (isopropyl m- chlorocarbanilate, 4.48 kg/ha), etho-
fumesate (2-ethoxy-2, 3-dihydro-3, 3-dimenthyl-5-benzofuranyl methane-
sulfonate, 2.24 kg/ha) or Stoddard oil (561 l/ha) applied to field
plots. Ethofumesate treatments to 'Scarlet Nantes' grown in the
greenhouse produced significant reductions in isolated periderm
weight per unit area. This decrease occurred both in the chloroform/
methanol-soluble and insoluble fractions. The diffusion resistance
of treated roots was significantly lower than controls. Thin layer
chromatography revealed no qualitative change in the composition of

the soluble extract.

Introduction

 

The outer surface of the mature carrot root is formed by peri-
derm, whose exterior row of cells may be isolated from underlying
tissue with the use of fungal pectinase. Due to the presence of the

suberin polymer (11) and associated soluble lipids (4), the periderm

31

32

cell walls, particularly the thickened, outer transverse wall, can
be compared to the typical cuticle of a leaf (for a comprehensive
description of cuticles see Martin and Juniper, 1970).

Cutin, another lipid polymer, forms the structural basis of
the cuticle, which typically contains waxes. These are often present
as surface fine structure as well as embedded in the cutin matrix.

The synthesis of cuticular waxes can be inhibited by herbi-
cides, most notably trichloroacetic acid (TCA) (3) and s-ethyl
dipropyl thiocarbamate (EPTC) (6), and significant reductions can
seriously affect the protective function of the membrane (2, 7, 13).
In the case of the thiocarbamate derivatives (9) the mode of action
is believed to be an inhibition of fatty acid elongation resulting
in reduced concentrations of other major wax components whose pre-
cursors are long chain acids (12).

Many of the embedded lipids of carrot periderm (4) are also
common to cuticular waxes (17). Carrot roots are probably dependent
upon the integrity of the periderm and the quantity of those soluble
lipids to control diffusion of substances into and out of the tissue.
Few studies on the effect of chemicals on suberin synthesis have been
published. Chlorpropham (isopropyl mychlorocarbanilate) was found
to inhibit wound periderm formation in potato at extremely low con-
centrations but did not prevent suberization of cells at the wound
surface (15). Soluble lipid synthesis in potato wound periderm was
decreased by TCA with the result that the loss of water vapor from

the tubers was increased (16).

33

This paper reports findings on the effect of foliar-applied

herbicides on carrot root periderm in field and greenhouse studies.

Materials and Methods

 

General.--All solvents were glass-distilled. Filter papers
and thin layer plates were prewashed in the solvent to be used with

them. Plates were activated at 100 C immediately prior to use.

Plant material and culture.--Seed of 'Danvers' and 'Scarlet

 

Nantes' were purchased from Joseph Harris Co., Rochester, New York.

A field trial of 5 herbicides on 'Danvers' was performed at
the Michigan State University Muck Soils Research Farm in Laingsburg,
Michigan. Seeds were sown in 15.2 m rows, 30 cm apart to form 1.2
x 7.6 m plots containing 3 rows. A randomized complete block design
with 4 blocks was utilized. Linuron (3-(3, 4-dichloropheny1)-l-
uethoxy-l-methylurea, 1.12 kg/ha), chlorpropham (isopropyl m-chloro-
carbanilate, 4.48 kg/ha), Stoddard oil (561 1/ha) and ethofumesate
(2-ethoxy-2, 3-dihydro-3, 3-dimethyl-5-benzofuranyl methanesulfonate,
2.24 kg/ha) were applied as foliar sprays 4 and 7 weeks after sowing
(Appendix D). All treatments except control received linuron (2.24
kg/ha) as a pre-emergence spray at the time of planting.

Based on the results of the field investigation, ethofumesate
was selected for a greenhouse experiment. Plants ('Scarlet Nantes')
were grown in vermiculite in lO-liter plastic pots and fertilized
weekly with half-strength modified Hoagland's (Appendix C) and Peter's
20-20-20 (rate = 200 ppm N). Five replicates of 3 pots were included

34

for each treatment. Ethofumesate (2.24 kg/ha) was applied as a
foliar spray when roots averaged 25 g fresh weight and harvest

followed 2 weeks later.

Lipid extraction and quantitation.--Roots harvested from

 

field plots (using only the middle row) were rinsed, dried and
extracted by immersing in a beaker of chloroform for l min. The
extracts were filtered, concentrated under reduced pressure and dried
under N2 for weighing. To determine soluble lipid weight/unit area,
the surface area of the roots was calculated. Assuming the roots
were a perfect circle in cross section, the following formula was
applied:
surface area = A, x i

where A1 is the area of the cut surface of a root sliced exactly down
the middle lengthwise. To determine this value, roots were laid cut-
side down on paper and their outline traced. These were cut out and
their area analyzed with an automatic area meter (Lambda Instruments).

Greenhouse-grown 'Scarlet Nantes' were harvested, rinsed and
the periderm was isolated and extracted. Briefly, peelings and cir-
cular discs were incubated for 4 days in 1% (w/v) buffered fungal
pectinase (pH 3.8). Following rinsing and air-drying, soluble lipids
were extracted. Approximately 200 to 300 mg periderm material were
refluxed twice for 2 hours with 25 m1 chloroform/methanol (CHCl3/Me0H
2:1 v/v). Total membrane weight, soluble lipid weight and insoluble

material were calculated on a unit area basis.

35

Thin layer chromatography.--Soluble lipids from ethofumesate-
treated and control plants of 'Scarlet Nantes' were dissolved in
chloroform to 250 cm2/ml, based on the calculation of their weight/
unit area. Two hundred p1 were applied in short bands to 0.25 mm
silica gel G plates. Development in benzene was followed by a 50%
sulfuric acid spary and charring at 100 C. Dissolved cabbage wax
(10 mg/ml) whose components had been previously documented (1, 5)

was also spotted for comparison.

Diffusion resistance measurements.--Diffusion resistance

 

(Rt) of control and ethofumesate-treated roots of 'Scarlet Nantes'
was determined by the method of Kolattukudy and Dean (10) using the

formula:

where P and Pa are vapor densities of the tissue and the air respec-

t
tively and E is the evaporation from the root in g/sec/cmz. The

surface area of the roots was determined as before.

Results and Discussion

Field tria1.--The results of the herbicide field experiment

 

are summarized in Table 3. No significant differences were evident
after statistical evaluation, even though a range of lipid concentra-
tions was found. The coefficient of variation in this experiment
was, unfortunately, 28%. This is a problem commonly encountered when

analyzing periderm lipids from field plots.

36

TABLE 3.--Effect of herbicides on the concentration of soluble peri-
derm lipids in 'Danvers'.2

 

 

T Rate Soluble lgpids
reatment (kg/ha) (pg/cm )
CIPC 4.48 94
Ethofumesate 2.24 89
Stoddard solvent 551y 108
Linuron 1.12 101
Control

Linuron pre-emerge 2.24 112

No pre-emerge -- 109

 

2Data are the average of 4 replicates.
yLiters/ha.

37

The method of lipid extraction (dipping the subject in a
beaker of solvent) and calculation of root surface area are consid-
erably less exact than the procedure using isolated membranes, but
is quite convenient when large numbers of roots need to be screened.
Although Espelie et a1 (4) maintain that this is a reasonably valid
method of lipid collection for qualitative analysis, a simple experi-
ment indicated to us that it may give an inaccurate indication of
periderm lipid concentration. A 2 x 2 factorial experiment using 2
solvents (CHCl3 and 2:1 CHClleeOH) and 2 lipid extraction methods
(immersion of root and refluxing isolated membrane) gave the results
shown in Table 4. Immersion in CHCl3/Me0H probably removes lipids
from internal tissue, and immersion in chloroform alone may not
completely extract all soluble lipids embedded within the suberin-
cell wall matrix (16). It was concluded that refluxing isolated mem-
branes in CHCl3/Me0H would result in the most complete extraction of

the compounds of interest without contamination from internal lipids.

Greenhouse trial of ethofumesate.--Ethofumesate gave the
lowest average value for soluble lipid concentration and therefore
was chosen for the greenhouse trial. The effect of the treatment
was to reduce both soluble and insoluble components of the periderm.
Table 5 illustrates the decrease in total membrane weight of plants
two weeks after receiving the herbicide. The decrease was due to
reductions in both soluble and lipid and the weight of the suberin-
enriched insoluble material. These results are consistent with those

of Leavitt et al who demonstrated inhibition of fatty acid metabolism

38

TABLE 4.--Comparison of periderm extraction techniques.Z

 

 

Weight ofy
extract
Solvent Method (pg/cmz)
CHCl3 Immersionx l4
Refluxw 23
CHCl3-MeOH Immersion 69
Reflux 43

 

2Data are the average of 4 replicates.

yF value for solvent significant at the 5% level.
x60 sec in chloroform. '

"Overnight in CHCl3/Me0H (2:1 v/v).

39

TABLE 5.--The effect of ethofumesate on the periderm composition of
'Scarlet Nantes'.z

 

Total membrane Soluble lipid Insoluble matrix
Treatment wt. (pg/cm?) wt. (pg/cm ) (pg/cm )

 

Ethofumesate 423*y 32* 400*
(2.24 kg/ha)

Control 481 42 429

 

2Data are the average of 5 replicates of 30-50 roots.

yF value significant at the 5% level.

40

by ethofumesate in cabbage (13). Fatty acids are components of both
carrot suberin (11) and its associated wax (4) and a block in their
synthesis or elongation could produce the results obtained here.
Flore and Bukovac (5) reported no change in 16- and lB-carbon
cutin acids with concentrations of EPTC that substantially inhibited
cuticular wax deposition in cabbage. Suberin alcohols, whose pre-
cursors are thought to be 16- and lB-carbon acids (8) are character-
ized by chain lengths longer than 18 and their synthesis may involve
an elongation system which is sensitive to ethofumesate. The coef-
ficient of variation in this experiment was less thatn 15% for all
parameters and significant differences were detected in this data as
noted in the table. The greenhouse environment and the more exact
analytical methods probably contributed to the reduction in variabi-

lity.

Qpalitative analysis.--Thin layer chromatography of soluble
lipids from both treatments in the greenhouse experiment revealed no
change in the relative amounts of the various wax components (Figure
8). An increase in long-chain esters resulting from inhibition of
fatty acid elongation was reported by Flore using EPTC (5) and
Leavitt with ethofumesate (13) but was not found in our study. The

former investigations used cabbage leaf wax as a experimental system.

 

Diffusion resistance.--The calculated Rt values for ethofume-
sate-treated and control roots of 'Scarlet Nantes' were 3.05 sec/cm

and 4.56 sec/cm respectively. These means are significantly different

41

0.75

0.40

0.28
0.23

 

Figure 8.--Thin lgyer chromatogram of soluble periderm lipids from
250 cm surface of 'Scarlet Nantes'. Developing solvent =

benzene. A. Control. B. Ethofumesate-treated (2.24 kg/ha).
C. Cabbage wax.

42

at the 5% level. Apparently the decrease in the soluble and/0r
insoluble periderm membrane fractions increased the permeability of
the periderm. This finding is consistent with that of Soliday (16),
using potato wound periderm, and other cuticle researchers who showed
that cuticular waxes are largely responsible for the impermeability

of the cuticle.

Summary and Conclusions

 

The periderm of the carrot root is vulnerable to the actions
of ethofumesate during its development as evidenced by a decrease
in soluble lipids and the insoluble portion of the membrane. The
implications of this herbicide effect were demonstrated by the
decreased diffusion resistance of the root tissue.~ A carrot suffer-
ing from the lipid synthesis inhibition reported here will probably
exhibit an increased rate of water loss when not in high humidity

storage conditions.

10.

43

Literature Cited

 

Baker, E. A. 1977. Sources of variability in plant epicuticu-
lar waxes and some effects on foliar penetration. Ph.D.
Thesis, University of Bristol.

Davis, D. G. and K. E. Dusbabek. 1973. Effect of diallate on
foliar uptake and translocation of herbicides in pea. Weed
Sci. 21: 16-18.

Dewey, D. R., P. Gregory, R. K. Pfeiffer. 1956. Factors affect-
ing the susceptibility of peas to selective dinitro herbicides.
Proc. 3rd. Br. Weed Contr. Conf. 1: 313-326.

Espelie, K. E., N. Z. Sadek, and P. E. Kolattukudy. 1980.
Composition of suberin-associated waxes from the subterranean
storage organs of seven plants. Planta 148: 468-476.

Flore, J. A. and M. J. Bukovac. 1978. Pesticide effects on
the plant cuticle: III. EPTC effects on the qualitative
composition of Brassaia oleracea L. leaf cuticle. J. Amer.
Soc. Hort. Sci. 103: 297-301.

Gentner, W. A. 1966. The influence of EPTC on external foliage
wax development. Needs 14: 27-31.

Juniper, B. E. 1957. The effect of pre-emergence treatment of
peas with trichloroacetic acid on the submicroscopic struc-
ture of the leaf surface. New Phytol. 58: 1-5.

 

Kolattukudy, P. E. 1978. Chemistry and biochemistry of the
aliphatic components of suberin. p. 43-83. In G. Kahl (ed.)
Biochemistry of wounded plant tissues. de Gruyter, New York.

Kolattukudy, P. E. and L. Brown. 1974. Inhibition of cuticular
lipid biosynthesis in Pisum sativum by thiocarbamates. Plant
Physiol. 53: 902-906.

 

Kolattukudy, P. E. and B. B. Dean. 1974. Structure, gas chroma-
tographic measurement and function of suberin synthesized by
potato tuber tissue slices. Plant Physiol. 54: 116-121.

11.

12.

13.

14.

15.

16.

17.

44

Kolattukudy, P. E., K. Kronman and A. J. Poulose. 1975. Deter-
mination of the structure and composition of suberin from
the roots of carrot, parsnip, rutabaga, turnip, red beet and
sweet potato by combined gas-liquid chromatography and mass
spectrometry. Plant Physiol. 55: 567-573.

 

Kolattukudy, P. E. and T. J. Walton. 1973. The biochemistry
of plant cuticular lipids. ProgyyChem Fats other Lipids.
13: 119-175.

 

Leavitt, J. R. C., D. N. Duncan, D. Penner and W. F. Meggit.
1978. Inhibition of epicuticular wax deposition on cabbage
by ethofumesate. Plant Physiol. 61: 1034-1036.

 

Martin, J. R., and B. E. Juniper. 1970. The cuticles of plants.
St. Martin's Press, Inc., New York.

Reeve, R. M., L. J. Forrester and C. E. Hendel. 1963. Histolo-
gical analysis of wound healing in potatoes treated in inhibit
sprouting I. CIPC (isopropyl-N-3-chlorophenyl carbamate)
treatments. J. Food Sci. 28: 649-654.

 

Soliday, C. L., P. E. Kolattukudy and R. W. Davis. 1979. Chemi-
cal and ultrastructural evidence that waxes associated with
the suberin polymer constitute the major diffusion barrier
to water vapor in potato tuber (Solanum tuberosum L.) Planta
166: 607-614.

 

Tulloch, A. P. 1976. Chemistry of waxes of higher plants.
p. 235-287. In P. E. Kolattukudy (ed.) Chemistry and bio-
chemistry of natural waxes. Elsevier Amsterdam, NY.

SECTION III
THE FUNCTION AND CHEMISTRY OF THE CARROT ROOT
PERIDERM IN LONG-TERM STORAGE
Abstract

Carrot roots (Daucus carota L.) lacking periderm or soluble

 

periderm lipids were found to lose 38 and 23% of their fresh weight
respectively in 6 days compared to controls which lost 7% at 4 C and
88% relative humidity. Weight losses of all treatments in saturated
air were negligible. Diffusion resistance of both treatments
decreased when compared to controls. The periderm membrane isolated
from 'Spartan Bonus' and 'Tito' was found to increase in weight/unit
area during 6 months at 0 C and 100% relative humidity. Chloroform/
methanol-soluble components decreased in 'Tito' but increased in

'5. Bonus'. The insoluble membrane fraction of '5. Bonus' was
depolymerized with LiAlH4 and the suberin diols obtained were seen
to increase dramatically during the first 3 months. The increase

in suberin and soluble lipids did not account for the total increase

in weight of the membrane, indicating the synthesis of an additional,

unidentified component(s).

Introduction

 

Preserving the quality of carrot roots depends upon reducing

desiccation after harvest. Optimum storage conditions have been

45

46

established by van den Berg and Lentz as 0 C and 98-100% relative
humidity (18). This environment successfully provided for the main-
tenance of firm texture, one of the most important factors is consumer
acceptance of fresh market carrots (1). In atmospheres which are
less than saturated, a 5-7% fresh weight loss occurs rapidly, causing
the commodity to become softened and represents an economic loss (10).
Weight loss of carrots in long-term storage is primarily due to
transpiration rather than respiration (18).

The periderm 0f the mature root, by virtue of its location
at the surface, is a primary consideration in the passage of water
into or out of the root. The cell walls of this tissue are suberin-
enriched (12) and contain soluble lipids, or waxes (6, 16). The
periderm, therefore, of carrot and other storage organs (e.g., potato,
beet and turnip) is similar in chemistry and structure to a cuticle,
but differs in certain aspects. The cuticle (reviewed thoroughly by
Martin and Juniper, 1970) is a superficial, cutinized membrane lying
on top of the epidermal tissue of a leaf, for example, where it
functions to reduce water loss. Although waxes impregnate the cutin
matrix, it is common for some of the cuticular lipids to be present
as surface fine structure (13). These epicuticular waxes are hydro-
phobic in character and have proven to be the most important wax com-
ponent with respect to limiting cuticle permeability (2, 3). The
soluble lipids associated with the suberized portions of the carrot
root periderm have been partially characterized and include alkanes,

esters, alcohols and fatty acids (6), compounds typical of cuticular

47

waxes (17). However, the carrot periderm lipids are relatively more
polar (6) than at least one extensively studied leaf wax: cabbage
(7).

The biosynthesis of periderm lipids in underground crops has
not been investigated, although suberin synthesis during wound heal-
ing has been described (4, 5). Additional information is available
on the function of the soluble lipids of potato wound suberin. These
compounds were shown to decrease the permeability of the periderm
(16). The qualitative composition as well as the quantity of waxes
may determine the effectiveness of the periderm as a diffusion barrier.
Individual classes of cuticular wax have been shown to limit the dif-
fusion of water to varying degrees (3, 8).

In the investigation which follows, changes in the chemical
composition of the suberized cells covering the carrot root surface
were monitored to determine whether the synthesis of suberin or $01-
uble lipid was occurring in cold storage. An additional study gave

evidence of the function of the periderm in preventing desiccation.

Materials and Methods
General.--Fungal pectinase was obtained from United States
Biochemical Corporation, Cleveland, Ohio and the bis(trimethylsilyl)
acetamide from Aldrich, Inc., Milwaukee, Wisconsin. Filter paper
and thin layer plates were pre-washed in the appropriate solvent

before use. All solvents were glass-distilled.

Desiccation studies.--The contribution of periderm components

to the maintenance of carrot turgidity was studied in a 2 x 3

48

factorial experiment with 5 replicates of 3 roots composing each
treatment. Roots were peeled as lightly as possible with a common
kitchen paring tool to remove suberized periderm cells, or immersed
for l min in a beaker of chloroform (CHC13) to extract soluble
periderm lipids. A third group was left intact to form the controls.
Two relative humidities (RH) were employed; those roots exposed to
the air in the storage room formed the 88% treatment (vapor pressure
deficit = 0.03) while a standard environment was achieved by holding
the carrots in perforated polyethylene bags. The experiment was
conducted at 4 C and the subjects were weighed daily for 6 days.
Diffusion resistance (Rt) of control and extracted roots was
determined by the method of Kolattukudy and Dean (11) using the

formula:

where Pt and Pa are vapor densities of the tissue and the air,

respectively and E is the evaporation from the root in g/sec/cmz.

The surface area of the roots was determined by the formula:
Surface area = A1 x i

where Ai is the two dimensional area of the cut surface of a root

which has been sliced in half lengthwise.

Storage experiments.--Seeds of 'Spartan Bonus' were purchased
from Harris, Rochester, New York and 'Tito' roots were a gift from
the carrot breeding program at Michigan State University. Both

varieties were grown muck soils using standard cultural procedures.

49

Two separate experiments monitored the composition of the
periderm during long-term storage. The variety for the first trial
was 'Tito' and for the second, '5. Bonus'. Three replicates were
held in plastic bags at 0 C and sampled at 0, 3 and 6 months. At
each sampling date, periderm weight and soluble lipid content were
determined. One hundred, 6-mm discs were excised from the surface
of several roots of each replicate. From these discs, periderm cells
were isolated from underlying tissue with 1% fungal pectinase in
citrate-phosphate buffer (pH 3.8). Four days of incubation were
sufficient to remove all cellular debris from the suberized portion
of the periderm leaving thin sheets. The discs were then rinsed,
air-dried and weighed.

In preparation for lipid extraction, approximately 20-30
roots were peeled and the peels treated with pectinase as described
above. After drying, the membranes were refluxed with chloroform/
methanol (CHCl3/MeOH 2:1 v/v) overnight and filtered through pre-
washed filter paper. The weight of the soluble lipids was calculated
by weighing the membranes before and after extraction.

2 root

These lipids were diluted with CHCl3 to reflect 250 cm
surface area/m1 and spotted on 0.25 mm silica gel G plates. Two
hundred pl were applied as a 0.5 cm band and the plate developed in
benzene. Visualization was accomplished with a 50% sulfuric acid
spray followed by charring at 100 C.

Approximately 200-300 mg of the insoluble, suberin-containing

residue following CHCl3/MeOH extraction were ground in a Wiley Mill

50

through a #40 mesh screen. Soxhlet extraction with CHC13/Me0H (2:1
v/v) for 2 hours removed remaining traces of soluble lipids. Depoly-
merization via hydrogenolysis with LiAlH4 proceeded according to the
method of Kolattukudy and Dean (11). Briefly, the ground periderm
material was refluxed overnight with excess LiAlH4 in 25 ml tetra-
hydrofuran. After acidifying with concentrated HCl, the reaction
mixture was extracted with CHCl3 (3 times, 100 ml). The combined
extracts were concentrated under reduced pressure. The suberin mono-
mers thus obtained were separated by preparative layer chromatography
on 0.5 mm silica gel G with hexane/ether/methanol (20:10:5 v/v/v).
After development plates were sprayed with 0.1% ethanolic 2'-7'-
dichlorofluorescein. Lipids were located under UV light and diols
were identified with the use of an authenticated potato suberin diol.
These were removed from the silica gel by washing with warm CHC13/
MeOH (2:1 v/v) into small tubes.

After evaporation of the solvent under N2 the dried residue
was heated with 0.5 ml bis(trimethylsilyl) acetamide at 90.C for
30 min. The resultant trimethylsilyl (TMS) derivatives were separated
by gas-liquid chromatography on a Packard model 7300 gas chromotograph
equipped with a flame ionization detector. The column was packed
with 3% SE-30 on Gas Chrom 0 (80-100 mesh). Operating temperature
was 220 C with a N2 flow rate of 25 m1/min. Peak areas of €16 and
C18:] diol were calculated by the product of height x width at I
height. Identity of the diols was confirmed by combined gas chroma-
tography/mass spectrometry with an HP-5985 system operating at 70 eV.

Spectra were determined at apices of peaks in the total ion scan.

51

Results and Discussion

Desiccation experiments.--The desiccation rates of carrot
roots at 4 C were dependent upon the relative humidity and the pre-
sence or absence of the periderm. When held at 88% RH, after 6 days
the peeled samples had lost 18% of their original fresh weight, over
4 times that lost by the controls (Figure 9b). The removal of the
soluble lipids from the root surface by CHCl3 extraction resulted
in a 23% loss, also significantly greater than the roots with peri-
derm intact. Even though the act of peeling the roots was damaging
to the cells, the data nevertheless implicate the periderm as an
important barrier to the diffusion of water, and a significant por-
tion of its effectiveness is due to the soluble lipid. The various
cuticle components also contribute to the protection of the internal
tissue. The waxes are the major barrier to the diffusion of water
(9, 15), and 2, 4-0 (3). Norris and Bukovac (14) suggested that the
orientation and continuity characteristics of embedded cuticular wax
may help determine the permeability of a cuticle, in addition to the
absolute wax quantity. Soliday et a1 (16) have demonstrated suberin-
associated waxes were essential to the Rt of wound periderm. Figure
9a shows the behavior of carrots in a saturated atmosphere. Note
that water loss is minimal from all treatments, although the desicca-
tion rate is slightly higher for extracted roots. Variations in
periderm weight and lipid content between cultivars or in response
to external factors should not seriously affect the storage life of

roots held in adequately high humidities over the short run. However,

52

 

 

 

 

 

EXTIACTED
1.54
II
a:
$1.00 PEELED
8. CONTROL
r— .
<o.54
O 4 - .L f a a
40+»
:7, PEELED
O
.J
h 350:
1:
52
u:
3 300
:1
an
I.“
E I“ 25 *1-
3 ti , txrucno
S
«:20:-
.—
«<
15 1-
1° " . cannon.
5 db
0 t i— : .L t f i
0 1 2 3 4 5 6 7

DAYS IN STORAGE

Figure 9.--Desiccati0n patterns of carrot roots with and without
periderm or soluble lipids. A. Storage in 100% RH.
8. Storage in 88% RH. Temperature = 4 C. Vertical
bars indicate standard errors.

53

under less than ideal conditions, it appears desirable to maximize
the soluble and insoluble portions of the periderm.

The diffusion resistance of extracted roots (Table 6) was
found to be 3.13 sec/cm while the average for intact roots was 6.16.
This convenient parameter allows us to assign a numerical value to
the permeability of a membrane covering a 3 dimensional surface.
Our results further confirm the role of the soluble lipids in inhi-

biting the diffusion of water vapor through the periderm.

Storage studies.--The periderm of both 'Tito' and '5. Bonus'

 

proved to be a dynamic system, undergoing apparent synthetic activi-
ties during the 6 months in storage. Date for periderm weights are
listed in Table 7. For each sampling date, the weight of the suberin-
containing residue (total periderm) was separated into CHCl3/Me0H -
soluble and insoluble fractions. The total periderm increased signi-
ficantly in both cultivars but not to the same degree. 'Tito' had a
gain of 26% and '5. Bonus', 83%. The insoluble material from the
isolated periderm increased in a similar manner; 31% and 80% for
'Tito' and '5. Bonus', respectively. The trend of the soluble lipids,
or waxes, was different for the two cultivars, increasing substan-
tially in '5. Bonus' but decreasing in 'Tito'. Thin layer chromato-
graphy of soluble lipids revealed no qualitative changes in their
composition (Figure 10).

The increase in the non-soluble portion of the periderm sug-
gested that the synthesis of suberin was occurring. The technique

of Walton and Kolattukudy (19) provided a sensitive and convenient

54

TABLE 6.--Diffusion resistance of carrot roots lacking periderm or
soluble periderm lipids.z

 

 

Treatment Rt (sec cm'1)
Control 6.16a~y
CHCl3-extracted 3.13b
Peeled 3.38b

 

2Data are average of 5 replicates.
yMean separation by Duncan's Multiple Range Test, 5% level.

55

TABLE 7.--Periderm composition of 'Tito' and '5. Bonus' in long-term
cold storage.2

 

 

 

Months Total Soluble Insoluble
in periderm wt. lipid wt. material
Cultivar storage (pg/cm ) (pg/cmz) (pg/cm )
'11 to ' 0 628ay 59a 569a
3 743b 45ab 698ab
6 860C 41b 891b
'5. Bonus' 0 460a 37a 423a
3 677b 60b 617b
6 8560 83c 773C

 

2Data are the average of 3 replicates.

yMean separation within columns cultivar by Duncan's Multiple Range
Test, 5% level.

56

0.80

0.37

0.26
0.17
0.09

0.00

 

Figure lO.--Thin layer chromatogram of soluble periderm lipids from 'S.
Bonus' following various storage periods. A. Cabbage leaf
wax. B. 0 mos. C. 3 mos. D. 6 mos.

57

method for assaying the suberin content of the periderm. Hydrogenoly-
sis of suberin with LiAlH4 releases the aliphatic components of the
polymer with the reduction of all carboxylic functions to hydroxyls.
Since w-hydroxy acids and dicarboxylic acids account for a combined
total of 55% of the aliphatic fraction of carrot suberin (12), the
quantitation of their common hydrogenolysis product, the a, w diol

was employed as an indicator of the suberin content of the periderm.
This method has been used reliably for an assay of suberization in
potato tissue (11). Specifically over 80% of the diol fraction of
carrot is composed of the C16 and Cl8zl monomers (12).

Gas-liquid chromatography of the trimethylsilyl ethers of
diols is illustrated in Figure 11. Identification of the peaks was
confirmed by mass spectrometry. The average pg diol/unit area are
listed in Table 8. The data imply that a dramatic increase in
suberin occurred during the first 3 months of storage. Between 3
and 6 months however, suberin deposition did not contribute to the
general increase in the weight of the membrane. The percent of the
total periderm weight attributed to the polymer decreased during
these 3 months, indicating that synthesis of other membrane components
is occurring. The additional soluble lipids which accumulated during
this period do not account entirely for the increase. The origin of

this additional mass remains unknown at this time.

Summary_and Conclusion
Carrots and other root crops are not known to wound held in

the manner of potato tubers, however the data here demonstrate that

58

 

 

RELATIVE DETECTOR RESPONSE

 

 

 

 

 

A L I l I A J
U

A
' 1 ' ' I v 1

1 ' 5 10

A
'

it

RETENTION TIME (min)

Figure ll.--Gas chromatogram of TMS ethers of suberin diols from
'S. Bonus'. Column = 3% SE-30. N2 flow rate = 25 m1/min.
Temperature = 220 C.

59

TABLE 8.--Suberin diol content of 'S. Bonus' periderm in long-term

 

 

storage.z
Months
in 010132’
storage (pg/cm I
0 6.11:1"
3 19.37b
6 14.23b

 

zData are the average of 3 replicates.

yCombined C16 and C18,] diol.
Mean separation by Duncan's Multiple Range test, 5% level.

60

suberin synthesis does occur in the harvested root, even at low

(0 C) temperatures. Although we have not investigated the permeabi-

lity of roots before and after storage, based on our desiccation data
we predict that the additional soluble lipid present at 6 months will
lend additional resistance to the loss of fresh weight. The suberin

polymer and its free waxes both play a role in limiting the diffusion

of water vapor.

10.

11.

61

Literature Cited

Arthey, V. D. 1975. Quality of horticultural products.
Butterworth and Co., Ltd., London. '

Bain, J. M. and D. McG. McBean. 1967. The structure of the
cuticular wax of prune plums and its influence as a water
barrier. Aust. J. Biol. Sci. 20: 895-900.

 

Baker, E. A. and M. J. Bukovac. 1971. Characterization of the
components of plant cuticles in relation to the penetration
of 2, 4-D. Ann. Appl. Biol. 67: 243-253.

Dean, B. B. and P. E. Kolattukudy. 1977. Biochemistry of
suberization. Plant Physiol. 59: 48-54.

 

Dean, B. B. and P. E. Kolattukudy. 1976. Synthesis of suberin
during wound-healing in jade leaves, tomato fruit and bean
pods. Plant Physiol. 58: 411-416.

 

Espelie, K. E., N. Z. Sadek, and P. E. Kolattukudy. 1980.
Composition of suberin-associated waxes from the subterranean
storage organs of seven plants. Planta 148: 468-476.

Flore, J. A. and M. J. Bukovac. 1978. Pesticide effects on the
plant cuticle III. EPTC effects on the qualitative composi-
tion of Brassaia oleracea L. leaf cuticle. J. Amer. Soc.
Hort. Sci. 103: 297-301.

 

Grncarevic, M. and F. Radler. 1967. The effect of wax compo-
nents of cuticular transpiration-model experiments. Planta
75: 23-27.

Haas, K. and J. Scthherr. 1979. Composition of soluble cuti-
cular lipids and water permeability of cuticular membranes
from Citrus leaves. Planta 146: 399-413.

Hardenburg, R. E. 1951. Further studies on moisture losses of
vegetables packaged in transparent films and their effect on
shelf life. Proc. Amer. Soc. Hort. Sci. 57: 277.

Kolattukudy, P. E. and B. B. Dean. 1974. Structure, gas
chromatographic measurement and function of suberin synthe-
sized by potato tuber tissue slices. Plant Physiol. 54:
116-121.

12.

13.

14.

15.

16.

I7.

18.

19.

62

Kolattukudy, P. E., K. Kronman and A. J. Poulose. 1975. Deter-
mination of the structure and composition of suberin from
the roots of carrot, parsnip, rutabaga, turnip, red beet and
sweet potato by combined gas-liquid chromatography and mass
spectrometry. Plant Physiol. 55: 567-573.

 

Martin, J. T. and B. E. Juniper. 1970. The Cuticles of plants.
St. Martin's, New York.

Norris, R. F. and M. J. Bukovac. 1968. Structure of the pear
leaf cuticle with special reference to cuticular penetration.
Amer. J. Bot. 55: 975-983.

 

Schdnherr, J. 1976. Water permeability of isolated cuticular
membranes: The effect of cuticular waxes on diffusion of
water. Planta 131: 159-164.

Soliday, C. L., P. E. Kolattukudy and R. W. Davis. 1979. Chemi-
cal and ultrastructural evidence that waxes associated with
the suberin polymer constitute the major diffusion barrier
to water vapor in potato tuber (Solanum tuberosum L.). Planta
166: 607-614.

Tulloch, A. P. 1976. Chemistry of waxes of higher plants. In
P. E. Kolattukudy (ed.) Chemistry and biochemistry of natural
waxes. Elsevier, Amsterdam, NY.

van den Berg, L. and C. P. Lentz. 1966. Effect of temperature,
relative humidity and atmospheric composition on changes in
quality of carrots during storage. Food Technol. 20: 104-
107.

Walton, T. J. and P. E. Kolattukudy. 1972. Determination of
the structures of cutin monomers by a novel depolymerization
procedure and combined gas chromatography and mass spectro-
metry. Biochemistry 11: 1885-1896.

APPENDICES

63

64

APPENDIX A

Desiccation of Five Carrot Lines

65

APPENDIX A

Desiccation of Five Carrot Lines

 

Roots grown in muck soil at Imlay City, Michigan were held
approximately 5 months in polyethylene bags at 0 C. Fresh weight
loss after 142 days was correlated with the average surface/volume
ratio of each of the 5 lines (Table 9). Linear correlation was signi-
ficant at the 10% level. Surface to volume ratio was the only para-
meter found to be a determinant of desiccation rate in these carrot
lines.

Roots grown in muck soil at Grant, Michigan were held at 5 C,
exposed to the air for 19 days. Surface/volume ratios were signifi-
cantly different for the 5 lines as was fresh weight loss on certain
days (Table 10). Desiccation could not be correlated with surface/

volume at any sampling date.

TABLE 9.--Desiccation of 5 carrot lines at 0 C.2

66

 

 

Line % fresh wt. losty surface/volumex
MSU 1414 B 2.17 1.64
MSU 1410 B 3.28 1.99
Tito 2.21 1.39
MSU 1389 B 3.15 2.02
A Bak 1.60 1.50

 

2Data are the average of 3 replicates of 3 roots.
F. value not significant.

yWeight lost in 142 days.

xCorrelation with fresh weight 10St significant at 10% level.

r = 0.83.

67

TABLE 10.--Desiccation of 5 carrot lines at 5 C.2

 

% fresh weight lostz
Days in storpge surface/
Line 3 7 ll 15 T9 volume

 

MSU 1414 B 1.52 5.91 ay 8.43 11.21 a 18.95 a 2.27 bc
Tito 1.36 4.03 ab 7.60 12.57 a 17.74 a 1.56 a
A Bak 1.67 7.07 ab 8.80 12.90 a 20.14 a 2.02 ab
M50 1389 B 2.54 8.54 b 11.59 15.49 ab 19.23 a 2.69 c
MSU 1410 B 1.68 8.05 b 13.77 18.45 ab 28.16 b 2.50 bc

 

r 0.35 0.61 0.24 0.47 0.39

 

zData are the average of 3 replicates.

yMean separation within columns by Duncan's Multiple Range Test, 5%
level. Included only where F value was significant.

xCorrelation between fresh weight loss and surface/volume.

68

APPENDIX B

The Effect of Herbicides on Red Beet and
Sugar Beet

69

APPENDIX B

The Effect of Herbicides on Red Beet and
Sugar Beet

 

Red beets were grown at the Michigan State University Horti-
cultural Research Center at East Lansing, Michigan. Herbicides were
applied to a randomized complete block design experiment. The pre-
emerge ethofumesate spray was applied 5 days after sowing and all
others 7 weeks later. Harvest followed ll days after spray treatments.
Membrane weights were determined as described in the text for carrot.
Desiccation rates were determined for sample of 3 beets per replicate
held at 5 C (Table ll).

Sugar beets were grown at the Michigan State University
Horticultural Research Center, East Lansing, Michigan. Herbicides
were applied to a randomized complete block design experiment. Pre-
emergence ethofumesate was sprayed 5 days after sowing and all others
7 and 14 weeks after sowing. Harvest followed 2 weeks after second
application of treatments. Desiccation rates of samples of 3 roots

per replicate were determined at room temperature (Table l2).

70

TABLE ll.--The effect of herbicides on periderm membrane weight and
desiccation in red beet.z

 

 

Desiccation
Rate Membrane geight rate
Treatment (kg/ha) (pg/cm ) (% fw lost/day)

Ethofumesate
(post-emergence) 1.12 740 2.75
Ethofumesate
(pre-emergence) 2.24 1000 3.00
Cycloate 3.36 980 3.50
Pyrazon 4.48 870 l.75
TCA 6.72 890 2.50
EPTC 3.36 880 2.75
Control lOlO 2.50

 

2Data are the average of 4 replicates.

71

TABLE l2.--Desiccation rates of herbicide-treated sugar beets.2

 

 

Rate Desiccation rate
Treatment (kg/ha) (% fw lost/hr)

Ethofumesate

(post emergence) l.l2 0.55
Ethofumesate

(pre-emergence) 2.24 0.45
Cycloate 3.36 0.83
Pyrazon 4.48 0.77
TCA 6.72 0.80
EPTC 3.36 1.07
Control -- 0.63

 

2Data are average of 4 replicates of 3 roots.

72

APPENDIX C
Modified Half-Strength Hoagland's Solution

73

APPENDIX C

Table 13.--Modified, half-strength Hoagland's Solution

 

 

 

 

Chemical g/l
Solution A
Ca(N03)2 295.0
Sequestrene NaFe (13% EDTA) 38.44
Solution B
KH2P04 34.25
KNO3 l26.65
MgSO4-7H20 62.50
ZnS04-7h20 0.056
MnSO4-H20 0.39l
00504-5H20 0.021
Boric acid (H3803) 0.725
MoO3-2H20 0.0005

l ml A + l ml B in 500 ml H20

 

74

APPENDIX D

Herbicide Data

75

 

 

cwuzoa mpnmppm: 0 a3»

mumgpcmucoo mpaow$wmpzsm u own

:09 Rom <uh Ezwcom zoo uwum ovumuuocopguwgp <uk
u oo~-om_ mace;

mzowcm> _Po Roo_ mzowgm> mcwpwon .mwpmppwpmwn sampogpma p=m>Pom cgmvoopm
maocwnmnnga . Azmv

mm<m a: mm mm cwsmgxa m-Fxcmgaumuogoquueuo:PEmum :ongxm
mugspxgawsnpquospme

pcoaso >83 om xogos P-_-Apxcmgaoeopguvn-e .mv-m eogacwo
mumco$pzmmcmgpme
P»:mgzmo~cmnum-_aspmewu

mcomwm um om cogcoz -m .m-ogcxswu-m .~1»xogumum mummmszmosum

Lacczmpm ow N 5668“ momsmaeauo_;p_»8628_e P>:66-m Beam
mpmsmngmu

Lawmampm um o ummzuom -mcoxmcopuauowgppzzamuz szpmnm mpmopuzu

can mum e ua~1ogopsu mpmpwcmngmuoxopguLa pzaognomw Emsaogaogopnu

cmcapuw$=cmz cowpmpssgom msmz munch wEmz pmuwswsu msmz coesou

 

.ngauum$=cms use cowgmpasgom .wgzumpucwso: mcwuwngmzuu.ep apamh

o x~ozmaa<

BIBLIOGRAPHY

76

10.

77

BIBLIOGRAPHY

Baker, E. A. l974. The influence of environment on leaf wax
development in Brassica oleracea var. gemifera. New Phytol.
73: 995-996.

Baker, E. A. and M. J. Bukovac. 1971. Characterization of the
components of plant cuticles in relation to the penetration
of 2, 4-0. Ann. Appl. Biol. 67: 243-253.

Brieskorn, L. H. and P. H. Brinnemann. l975. Carbonsauren und
alkanole des cutins und suberins von Solanum tuberosum.
Phytochemistry 14: l363-l367.

 

 

Clarkson, 0. T. and A. H. Robards. l975. The endodermis, its
structure, development and physiological role. p. 415-436.
In J. G. Torrey and D. T. Clarkson (eds.) The development
and function of roots. Academic Press, New York.

Davis, D. G. and K. E. Dusbabek. l973. Effect of diallate on
foliar uptake and translocation of herbicides in pea. Heed
§g1_ 2l: l6-lB.

Dean, B. B. and P. E. Kolattukudy. l977. Biochemistry of suberi-
zation. Plant Physiol. Sl: 48-54.

 

Dean, B. B. and P. E. Kolattukudy. 1976. Synthesis of suberin
during wound-healing in jade leaves, tomato fruit and bean
pods. Plant Physiol. 58: 4ll-416.

 

Dean, B. B., P. E. Kolattukudy and R. W. Davis. 1977. Chemical
composition and ultrastructure of suberin from hollow heart
tissue of potato tubers (Solanum tuberosum) Plant Physiol.
59: lOOB-lOTO.

Dewey, 0. R., P. Gregory and R. K. Pfeiffer. 1956. Factors
affecting the susceptibility of peas to selective dinitro
herbicides. Proc. 3rd. Br. Heed Contr. Conf. l: 3l3-326.

Esau, K. l977. Anatomy of seed plants, 2nd ed. John Wiley and
Sons, Inc., New York.

ll.

12.

l3.

14.

15.

l6.

l7.

l8.

19.

20.

21.

22.

78

Espelie, K. E. and P. E. Kolattukudy. l979. Composition of
the aliphatic components of suberin of the endodermal fraction
from the first internode of etiolated Sorghum seedlings. Plant
Physiol. 63: 433-435.

Espelie, K. E., N. Z. Sadek and P. E. Kolattukudy. 1980. Com-
position of suberin-associated waxes from the subterranean
storage organs of seven plants. Planta 148: 468-476.

Flore, J. A. and M. J. Bukovac. l978. Pesticide effects on the
plant cuticle: III. EPTC effects on the qualitative composi-
tion of Brassica oleracea L. leaf cuticle. J. Amer. Soc.
Hort. Sci. ‘103: 297-301.

 

 

Grncarevic, M. and F. Radler. l967. The effect of wax compo-
nents of cuticular transpiration-model experiments. Planta
75: 23-27.

Haas, K. and J. Schbnherr. l979. Composition of soluble cuti-
cular lipids and water permeability of cuticular membranes
from Citrus leaves. Planta l46: 399-403.

Juniper, B. E. l957. The effect of pre-emergence treatment of
peas with trichloroacetic acid on the submicroscopic structure
of the leaf surface. New Phytol. 58: l-5.

 

Karunen, P. and L. Eronen. 1979. Influence of S-ethyl dipropyl-
thiocarbamate (EPTC) on the fatty acid composition of wheat
leaf galactolipids. Physiol. Plant 40: lOl-l04.

 

Kolattukudy, P. E. 1980. Biopolyester membranes of plants:
Cutin and suberin. Science 208: 990-l000.

Kolattukudy, P. E. 1978. Chemistry and biochemistry of the
aliphatic components of suberin. p. 43-84. In G. Kahl (ed.)
Biochemistry of wounded plant tissues. de Gruyter and Co.,
New York.

Kolattukudy, P. E. l977. Lipid polymers and associated phenols.
p. 185-257. In F. Loewus and V. C. Runeckles (eds.) Recent
Advances in Phytochemistry Vol. ll. Plenum Press, N.Y.

Kolattukudy, P. E. and V. P. Agrawal. 1974. Structure and com-
position of the aliphatic constituents of potato suber skin
(suberin). Lipids 9: 682-69l.

Kolattukudy, P. E. and L. Brown. l974. Inhibition of cuticular
lipid biosynthesis in Pisum sativum by thiocarbamates. Plant
Physiol. 53: 902-906.

 

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

79

Kolattukudy, P. E., R. Croteau and J. S. Buckner. l976. p. 290-
334. Biochemistry of plant waxes. In P. E. Kolattukudy (ed.)
Chemistry and biochemistry of Natural waxes. Elsevier,
Amsterdam, NY.

Kolattukudy, P. E. and B. B. Dean. l974. Structure, gas chroma-
tographic measurement and function of suberin synthesized by
potato tuber tissue slices. Plant Physiol. 54: ll6-l21.

Kolattukudy, P. E., K. Kronman and A. J. Poulose. l975. Deter-
mination of the structure and composition of suberin from the
roots of carrot, parsnip, rutabaga, turnip, red beet and sweet
potato by combined gas-liquid chromatography and mass spectro-
metry. Plant Physiol. 55: 567-573.

 

Kolattukudy, P. E. and T. J. Walton. 1973. The biochemistry of
plant cuticular lipids. Prog. Chem. Fats other Lipids. l3:
llQ-l75.

Laetsch, W. M. l974. The C4 syndrome. A structural analysis.
Ann. Rev. Plant Physiol. 25: 27-52.

Leavitt, J. R. C., D. N. Duncan, 0. Penner and N. F. Meggit.
l978. Inhibition of epicuticular wax deposition on cabbage
by ethofumesate. Plant Physiol. 6l: 1034-1036.

Leece, D. R. and A. L. Kenworthy. 1972. Influence of epicuticu-
lar waxes on foliar absorption of nitrate ions by apricot leaf
disks. Aust. J. Biol. Sci. 25: 64l-643.

 

Luttge, U. l97l. Structure and function of plant glands.
Ann. Rev. Plant Physiol. 22: 23-44.

Norris, R. F. l974. Penetration of 2,4-D in relation to cuticle
thickness. Am. J. Bot. 6l: 74-79.

 

Norris, R. F. and M. J. Bukovac. 1972. Influence of cuticular
waxes on penetration of pear leaf cuticle by l-NAA. Pesti.
Sci. 3: 705-708.

Macey, M. J. K. 1970. The effect of light on wax synthesis in
leaves of Brassica oleracea. Pbytochemistry 9: 757-76l.

 

Martin, J. T. and B. E. Juniper. l970. The cuticles of plants.
St. Martin's Press, Inc., New York.

Peterson, C. A., R. L. Peterson and A. N. Robards. l978. A
correlated histochemical and ultrastructural study of the
epidermis and hypodermis of onion roots. Protoplasma 96:
l-2l.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

80

Possingham, J. V., T. C. Chambers, F. Radler and M. Grncarevic.
1967. Cuticular transpiration and wax structure and composi-
tion of leaves and fruit of Vitis vinifera. Aust. J. Biol.
Sci. 20: ll49-llS3.

 

Reeve, R. M., L. J. Forrester and C. E. Hendel. I963. Histolo-
gical analysis of wound healing in potatoes treated to inhibit
sprouting I. CIPC (isopropyl-N-3-chlorophenyl carbamate)
treatments. J. Food Sci. 28: 649-654.

 

Riley, R. and P. E. Kolattukudy. 1975. Evidence of covalently
attached coumaric and ferulic acids in cutins and suberins.
Plant Physiol. 56: 650-654.

 

Schnepf, E. l974. Gland cells. p. 33l-357. In A. N. Robards
(ed.) Dynamic aspects of plant ultrastructure. McGraw-Hill,
New York.

Schfinherr, J. l976. Water permeability of isolate cuticular
membranes: The effect of cuticular waxes on diffusion of
water. Planta l3l: lS9-l64.

Scott, F. M. 1950. Internal suberization of tissues. Bot. Gaz.
111: 378-394.

Soliday, C. L., P. E. Kolattukudy and R. W. Davis. 1979.
Chemical and ultrastructural evidence that waxes associated
with the suberin polymer constitute the major diffusion
barrier to water vapor in potato tuber (Solanum tuberosum
L.). Planta l66: 207-2l4.

Still, G. 6., D. G. Davis and G. L. Zander. 1970. Plant epi-
cuticular lipids: Alteration by herbicidal carbamates.
Plant Physiol. 46: 307-314.

 

Nhitecross, M. I. and D. J. Armstrong. l972. Environmental
effects on epicuticular waxes of Brassica napus L. Aust. J.
Bot. 20: 87-95.

Wilkerson, R. E. and u. S. Hardcastle. l970. EPTC effects on
total leaflet fatty acids and hydrocarbons. Need Sci. 18:
125-128.