ANATOMICAL STUDY OF THE ANTHURIUM PLANT,
ANTHURIUM ANDREANUM, L., AND A COLOR
BREAKDOWN DISORDER OF ITS FLOWER

Dissertation for the Degree of Ph. D.
MICHIGAN STATE UNIVERSITY
TADASHI HIGAKI
1976

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[VIICI‘E . , .230
University

 

This is to certify that the

thesis entitled “ "4"" ' u. '

Anatom1ca1 Study of the Anthurium PTant,
Anthurium Andreanum, L. , and a Color
Breakdown Disorder of-its FTower

 

presented by

Tadashi Higaki

has been accepted towards fulfillment
of the requirements for

Ph.D. degree in HorticuIture

 

     
 

Q/C/ i Maw/94M"

Major fessor

 

./ r

Date February 22, 1977

 

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II.

 

 

ABSTRACT

ANATOMICAL STUDY OF THE ANTHURIUM PLANT,
ANTHURIUM ANDREANUM, L., AND A COLOR
BREAKDOWN DISORDER OF ITS FLOWER

I. Anatomical study of the anthurium plant,
Anthurium andreanum, L.

 

II. Color breadkown in anthurium flowers,
Anthurium andreanum, L.

 

BY
Tadashi Higaki

Section I

 

The gross morphology and anatomy study on Anthurium

 

andreanum, L., was by whole plant observation, using

 

dissecting and light microsc0pe, and scanning electron
microscopy. Anthurium is a perennial-herbaceous monocoty-
ledon in the family Araceae. It is low growing with
chordate leaves and attractive chordate flowers. It has a
juvenile phase when each leaf axil has a lateral vegeta-
tive bud and a generative phase when each leaf axil has a
flower and the lateral vegetative bud is located opposite
the leaf attachment. The "commercial flower” consists of a
conspicious bract (spathe) and a protruding rachis (spadix).
Minute, botanically perfect flowers are borne spirally on

the spadix. The flowers are protogynous as the stigma is

receptive <
the spathe
with l or 1
and lower 1
cells. Va:
the spathe
hypodermal
to the spa
ers of pal
below the

the mesopr
pedicel, E
Outer epid
SClerifiec
bundles We
were Cylir
were Chara
cells Cal]
found Scat

ground Par

Sec '
w

A €010
an

%.

SEctiOn

Tadashi Higaki

receptive one week before shedding of pollen. Anatomically,
the spathe has a one cell layered upper and lower epidermis,
with l or 2 layers of hypodermis cells. Between the upper
and lower hypodermis are 10-12 layers of spongy parenchyma
cells. Vascular bundles are dispersed uniformly throughout
the spathe. Anthocyanin pigments are localized in the
hypodermal cells. The leaf blade is similar in structure

to the spathe, except there is no hypodermis, but two lay-
ers of palisade parenchyma cells form the tissue immediately
below the epidermis. Cholorplasts were dispersed throughout
the mesophyll, but concentrated in the palisade cells. The
pedicel, petiole and vegetative stem are typically monocot.
Outer epidermal cells covered the cortex, a layer of
sclerified parenchyma cells and the ground tissue. Vascular
bundles were dispersed throughout the ground tissue. Roots
were cylindrical, fleshy, epiphytic and adventitious. They
were characterized by having multiple layers of epidermal
cells called the velamen. Raphide and druse crystals were
found scattered throughout the entire plant tissue. Above

ground parts were covered with a thick waxy layer of cuticle.

Section II

 

A color breakdown disorder:h1the spathe of Anthurium

 

andreanum, L., was investigated. Ca deficiency in the lobe

 

section of the spathe was found to cause the disorder.
Elemental analysis of spathe and leaf tissue of color break-
down andnormal plants revealed lower Ca in color breakdown

plants (color breakdown - spathe: 0.372%, leaves: 0.363%;

_.._—_
*
.—~
_——

..__

.—

—

H

_

.—

a

normal - Spat:
probe x-ray a
section of tt
mesophyll tis
breakdown syr
Symptoms wen
heads on the
The dots mul'
coalesced to
dehydrated a
0f complete
down in the
spathe and 1
4' higher at
in the lobe
regardless c
anall/Sis prg
lower uptakI
that calciu
field tESt
application
diSordeI.

spathe is s
critieal 1;
tissue (306;1
may be med

and temper.

 

Tadashi Higaki
normal - spathe: 0.830%, leaves: 0.805%). Electron micro—
probe X-ray analysis revealed lower Ca in lobe than tip
section of the spathe and higher Ca in epidermal than
meSOphyll tissue. Nutrient culture studies produced color
breakdown symptoms in the spathe with No Ca treatment.
Symptoms were tiny water-soaked lesions the size of pin
heads on the upper epidermal surface of the spathe lobe.

The dots multipled, increased in size and eventually
coalesced to form large water-soaked lesions. The lesions
dehydrated and turned brown. pH levels 3, 4, 5, 6, and 7

of complete nutrient solutions did not produce color break-
down in the spathe, although elemental chemical analysis of
spathe and leaf tissues indicated lower Ca uptake of 3 and
4, higher at 6, and highest at pH 7. Lower Ca was found

in the lobe than tip sections of both spathe and leaves,
regardless of treatment. In a separate study, X-ray
analysis produced similar results. In addition pH 9 gave
lower uptake of Ca than 7. Microautoradiography study showed
that calcium was deposited primarily in the cell wall. A
field test with different Ca sources confirmed that the
application of Ca significantly reduced the incidence of the
disorder. A critical level of Ca within the tissue of the
spathe is suggested with respect to the disorder. The
~critica1 level may be very narrow, but must be exceeded if
tissue does not exhibit color breakdown. The critical level
may be mediated by environmental conditions such as humidity

and temperature. Like oedema disorder which is correlated

fl—_—

‘

with environr‘
transpiratior
breakdown ma
Unlike oedem
lobe of the

color breakd
breakdown an

cells in aff

on the perma
lamella due

intumescence

Tadashi Higaki
with environmental conditions which hinder the normal
transpiration processes of the plant, anthurium color-
breakdown may similarly be related to transpiration.

Unlike oedema, however, the critical level of Ca in the

lobe of the spathe is the major factor in the anthurium
color breakdown disorder. Anatomical studies of color-
breakdown and normal flowers showed collapsing mesophyll
cells in affected spathe. Separation of cells, when present
on the permanent slides, suggested breakdown of the middle
lamella due to lack of Ca. No sign of hypertrophy or

intumescence was found.

 

ANAT OF

ANT}

ANATOMICAL STUDY OF THE ANTHURIUM PLANT,
ANTHURIUM ANDREANUM, L., AND A COLOR

BREAKDOWN DISORDER OF ITS FLOWER

BY

Tadashi Higaki

A DISSERTATION

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

Department of Horticulture

1976

‘w
m.
PF.

M

h

_

~—

A

A

———.—

 

I woul

all those V
studies, a:
To my

my heartfe
COuragemen
To tr
Carpenter‘
MaXine S.
haVe beerr
To m
Hawaii HO
appreCiat
ducive tc
To 1

and eXPE:
Bulleck'

Kunisaki

ACKNOWLEDGEMENTS

I would like to express my sincere appreciation to
all those who have helped me during the course of these
studies, as well as throughout my graduate education.

To my major professor, Dr. H. Paul Rasmussen, goes
my heartfelt thanks for his excellent counsel and en-
couragement.

To the members of my guidance committee: Drs. W. J.
Carpenter, A. L. Kenworthy, W. H. Carlson, M. E. Miller, and
Maxine S. Ferris, I am most grateful. Your help and advice
have been most gratifying.

To members of the Michigan State and University of
Hawaii Horticulture Departments, I would like to express my
appreciation for providing an excellent environment con-
ducive to such a meaningful graduate experience.

To the following individuals for their timely assistance
and expert advice: Esther M. Higaki, Michiko Higaki, Drs. R.
Bullock, Y. Sagawa, F. Laemmlen, Mr. V. Shull, and Mr. J.
Kunisaki, my humble thanks.

Appreciation is also extended to Mr. Harold T. Tanouye
of Hawaiian Anthuriums, Inc., for supplying anthurium flowers

and shipping them to Michigan.

ii

 

 

To ti
National I
of my fel

My 5
being a t

Fina
Pauline a

tolerance

To the Farm Foundation Scholarship Program and the
National Defense Act, I am most indebted for the funding
of my fellowship.

My special appreciation to Mr. Gordon Shigeura for
being a true friend especially when needed.

Finally, to my wife, Jean, and children Joanne, Connie,
Pauline and Chad for their encouragement, inspiration and

tolerance, my deepest heartfelt thanks.

iii

a
h
A
/——‘

'—
,_.._

 

 

 

LIST OF TA
LIST OF FI
INTRODUCTI

LITERATURE

Literature
Materials
RESultS a!

TABLE OF CONTENTS

 

 

 

Page

LIST OF TABLES . . . . . . . . . . . . . . . . . . v
LIST OF FIGURES . . . . . . . . . . . . . . . . . vi
INTRODUCTION . . . . . . . . . . . . . . . . . . . 1
LITERATURE CITED . . . . . . . . . . . . . . . . . 5

SECTION I

ANATOMICAL STUDY OF THE ANTHURIUM PLANT,
ANTHURIUM ANDREANUM, L.

Literature Review . . . . . . . . . . . . . . . . 10
Materials and Methods . . . . . . . . . . . . . . 12
Results and Discussion . . . . . . . . . . . . . . 19
Literature Cited . . . . . . . . . . . . . . . . . 58

SECTION II

COLOR-BREAKDOWN IN ANTHURIUM FLOWERS,
ANTHURIUM ANDREANUM, L.

Literature Review . . . . . . . . . . . . . . . . 61
Materials and Methods . . . . . . . . . . . . . . 67
Results and Discussion . . . . . . . . . . . . . . 82
Literature Cited . . . . . . . . ... . . . . . . . 98

APPENDIX
Correlation of the anatomical study and the
spathe color-breakdown disorder of
Anthurium andreanum, L. . . . . . . . . . . . . . 103
Literature Cited . . . . . . . . . . . . . . . . . lll

iv

— __————
~
_‘
,———_
“Wk—K
___——_—
—_‘
5—.“

 

 

 

Table

Solut:
and C1

Eleme
and 1
Plant

Ca x-
Spath
Color

Ca CC
leaf
and ;

Field
breap
anthr

LI ST OF TABLES
Table Page
SECTION II

1. Solution culture composition, concentration
and conditions used in the Ca experiment . . . . 77

2. Elemental analysis of anthurium spathe
and leaf from color-breakdown and normal
plants . . . . . . . . . . . . . . . . . . . . . 84

3. Ca x-ray analysis of a cross-section of the
spathe of anthurium in normal and
color-breakdown flowers. . . . . . . . . . . . . 88

4. Ca content of anthurium spathe and
leaf tiSsues grown at various nutrient
and pH levels . . . . . . . . . . . . . . . . . 90

5. Field tests of the incidence of color-
breakdown disorder of spathe in
anthurium with different Ca sources . . . . . . 95

Figur

lo,

11.

12.

13.

14.

LIST OF FIGURES

 

 

Figure
SECTION I
1. An apparatus used for tissuemat
infiltration . . . . . . . . . . . . . . .
2. A mature anthurium plant, Anthurium
andreanum, L., cv. Red Ozaki . . . . . . .
3. Scanning electron micrograph of
rhombohedral crystals in the anthurium . .
4. A scanning electron micrograph of the
upper cuticle of anthurium leaf . . . . .
5. The anatomy and morphology of the
anthurium flower . . . . . . . . . . . . .
6. Photomicrograph of the stamen of anthurium
7. Photomicrograph of the gynoecium of
anthurium . . . . . . . . . . . . . . . .
8. The various spathe shapes of the
anthurium flower . . . . . . . . . . . . .
9. Photomicrograph and SEM micrograph of
the spathe of the anthurium . . . . . . .
10. Photomicrographs and SEM micrographs
of the anthurium pedicel . . . . . . . . .
11. SEM and light photomicrographs of the leaf
of the anthurium . . . . . . . . . . . . .
12. Photomicrograph and SEM micrograph
of the anthurium root . . . . . . . . . .
13. Diagram of the vegetative stem of
the anthurium . . . . . . . . . . . . . .
14. Photomicrograph of the vegetative

meristem of the anthurium . . . . . . . .

vi

Page

16

21

25

27

30
32

35

38

41

43

46

50

53

56

_ __ .—
_~ _l_.L-—..-——— ._ M
”—u—n‘n ”W...
W

 

 

 

LIST OF

Figure

1. Ant
sta

2. Ill
lob
spa

3. Pla
in
rad

Mic
of
Eng

ant
gro

Spa

Col
SeP

LIST OF FIGURES-~Continued
Figure Page
SECTION II

1. Anthurium flowers showing various
stages of the color breakdown disorder . . . . 63

2. Illustration of sampling of tissue from
lobe and tip sections of the anthurium
spathe . . . . . . . . . . . . . . . . . . . . 73

3. Planting system used to grow anthuriums
in nutrient culture . . . . . . . . . . . . . 76

4. Diagram of apparatus used for
radioisotope study . . . . . . . . . . . . . . 80

5. Microprobe line profile x-ray analysis
of Ca in the spathe of anthurium plants . . . 86

6. Energy dispersive x-ray analysis of
anthurium flower sections of plants
grown at various nutrient and pH levels . . . 92

APPENDIX

1. A color breakdown and a normal anthurium
spathe tissue . . . . . . . . . . . . . . . . 106

2. Color breakdown spathe showing cellular
separation . . . . . . . . . . . . . . . . . . 109

vii

INTRODUCTION

The tropical anthurium, Anthurium andreanum, Lind., is

 

a member of the family Araceae, which includes more than
100 genera and 1,500 species. In Hawaii they are grown in
shade and high humidity. It is a perennial-herbaceous
plant cultivated for its attractive, long lasting flowers.
The flower is a complex of the colorful modified leaf
(spathe) and hundreds of small true flowers on the pencil-
1ike protrusion (Spadix) rising from the base of the spathe.

The anthurium plant produces flowers continuously
throughout the year. One flower emerges from each leaf
axil. The sequence of leaf, flower and new leaf is main-
tained throughout the life of the plant. The intervals
between leaf emergence depend on the natural changes in
environmental conditions. More flowers are produced during
the summer months, when conditions are favorable for growth
than during the winter months when temperatures are lower
and light intensity is reduced (26).

The anthurium, a native of Central America, was brought
to Hawaii from London in 1889 (27). Today, after 87 years
of cultivation and hybridization in Hawaii, the Hawaiian
anthurium flower is one of the State's principal ornamental

exports to the mainland U.S., Canada, Australia, Japan,

2

Germany, Holland, Italy and many other countries. Anthurium
culture has developed from a hobby or backyard operation 10
years ago, into a large-scale commercial enterprise, valued
at approximately 2 million dollars at the farm level (8).
The Hawaii State General Agricultural Plan (7) projects an
industry valued at 16.2 million dollars by 1989. Its future
for economic growth appears unlimited. The variety of
colors and the exotic beauty of the flower have attracted
many consumers and the demand for the Hawaiian anthurium is
steadily increasing. The long shelf-life and ease in han-
dling and packaging make it a durable product for shipping
and mailing throughout the world. The Hawaii State legis-
lature and the County government in Hawaii are increasingly
supporting market promotion and develOpment; farm loan
programs; and research programs on production, post harvest,
engineering and marketing problems of anthuriums. The
future looks bright and growth seems inevitable.

Total acreage in anthurium production in the State is
320 acres, of which 95% is on the Island of Hawaii (8). The
major acreage is confined to the windward portion of the
island where the weather is ideal with cool temperatures
(70-80°F day; 60-70°F night temperature) with abundant
rainfall throughout the year (170-200 inches annually) with
relative humidities of 75-100%.

This investigation examines the gross anatomy of the

plant and a disorder of the spathe of Anthurium andreanum,

 

 

Lind.

3

Information on the anatomy of Anthurium andreanum is

 

limited. Yet, the need to know thoroughly the nature and
function of the plant is the foundation for solving produc—
tion and handling problems. Realistic interpretations and
knowledge of the plant are imperative if Hawaii's anthurium
research program is to develop properly and efficiently.
This study examines extensively the external and internal
anatomy and morphology of the anthurium plant. Established
techniques in histochemistry and electron microscopy (31)
were used to accomplish the objectives.

The second section of this dissertation deals with color

and tissue breakdown in spathe of Anthurium andreanum, Lind.

 

Much of the limited research on Anthurium andreanum has

 

been done in Hawaii including studies on keeping quality
(1, 17, 32, 33, and 34), cost of production (6, 13, 14, and
15), cultural problems (2, 3, 5, 9, 10, ll, 12, 16, 18, 21,
25, 26, 28, 29, and 30), and breeding (4, 19, 20, 22, 23,
and 24). Except for long range breeding, most research is
problem-solving as it becomes acute in the industry. One
problem facing the industry was a flower color and tissue
disorder. The growers were faced with heavy losses due to
a disorder that caused water-soaked lesions and color-
breakdown on the lobes of the flower spathe. These water-
soaked lesions, in time dessicated, turned dark brown and
eventually dried. The flowers were unfit for sale. Field
losses of up to 50% have been reported and losses after

shipments to the overseas market of up to 20%. Many times

4

the visual symptoms were not evident during grading and
packing, but were manifest after transit to the distant
market. The following objectives were established for this
study.

1. A well-balanced training in agricultural

research, practical as well as basic.
2. A contribution of basic information on

Anthurium andreanum, Lind., that could be

 

utilized worldwide.
3. Solving of an acute problem on anthurium

production for Hawaii's agriculture.

LITERATURE C ITED

10.

11.

12.

13.

LITERATURE CITED

Akamine, E. K. and T. Goo. 1972. Relationships
between phsyical characteristics and vase life of
anthuriums. Haw. Farm Sci. 21(2): 9-10.

Aragaki, M. 1964. Maneb will control anthurium spadix
rot. Haw. Farm Sci. 13(4): 16.

and M. Ishii. 1960. A spadix rot of anthurium
in Hawaii. Plant Dis. Rep. 44(11): 865-867.

, H. Kamemoto and K. M. Maeda. 1968.
Anthracnose resistance in anthuriums. Haw. Agri. Expt.
Sta. Prog. Report No. 169.

, M. Sherman and H. Kamemoto. 1957. Phytotox-
icity of chemical sprays to anthuriums. Haw. Farm Sci.
5(3): 10-11.

Barmettler, R. and J. Sheehan. 1963. A budgetary
analysis for large-scale anthurium Operation in Hawaii.
Haw. Agri. Expt. Sta. Ag. Econ. Report No. 63.

Department of Planning and Economic Development. 1970.
Opportunities for Hawaiian agriculture. Ag. Dev. Plan.
State of Hawaii, Honolulu, Hawaii.

Hawaii Department of Agriculture. 1974. Statistics of
Hawaiian agriculture. Hawaii Crop and Livestock
Reporting Service, Honolulu, Hawaii.

Hayward, A. C. 1972. A bacterial disease of anthurium
in Hawaii. Plant. Dis. Rep. 56(10): 904-908.

Higaki, T. 1973. Chemical weed control in anthurium.
Haw. Agri. Expt. Sta. Res. Rep. No. 212.

and D. P. Watson, 1967. Anthurium culture in
Hawaii. Haw. Coop. Ext. Ser. Cir. No. 420.

and M. Aragaki. 1972. Benomyl for control of
anthurium anthracnose. Haw. Farm Sci. 21(3): 7-8.

Hiroshige, H. H. and T. Shirakawa. 1964. Anthurium
cost survey in Hawaii. U of H Farm Mangt. Series No. H1.

14.

15.

16.

17.

 

18.

19.

20.

21.

22.

23,

24.

DCGS

NEH

27.

 

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

7

and T. Higaki. 1970. Anthurium cost of pro-
duction survey under Saran in Pahoa, Hilo, and Mountain
View areas. U of H Farm Mangt. Series No. H8.

, and G. Aoki. 1966. Anthurium cost
survey under Saran in Hawaii. U of H Farm Mangt.
Series No. H6.

Hunter, J. E., W. H. Ko, R. K. Kunimoto and T. Higaki.
1974. Foliar nematode disease of anthurium seedlings.
Phytopath. 64(2): 267—268.

Kamemoto, H. 1962. Some factors affecting the keeping
quality of anthurium flowers. Haw. Farm Sci.
11(4): 2-5.

and H. Y. Nakasone. 1953. Effects of media on
prSduction of anthuriums. Haw. Agri. Expt. Sta. Prog.
Notes No. 94.

and . 1955. Improving anthuriums
through breeding. Haw. Farm Sci. 3(3): 4-5.

and . 1963. Evaluation and improvement
of anthurium clones. Haw. Agri. Expt. Sta. Tech. Bul.
No. 58.

and . 1957. WOod shavings as a medium
for anthuriums. Haw. Agri. Expt. Sta. Circ. No. 53.

and and M. Aragaki. 1968. Improvement
of anthuriums through breeding. Proc. Amer. Soc. Hort.
Sci. Trop. Region. 12: 267-273.

, Mr. Aragaki, T. Higaki and J. T. Kunisaki.
1969. Three new anthurium cultivars. Haw. Farm Sci.
18: 1-3.

, , and . 1971. A new
anthracnose resistant white anthurium. Haw. Farm Sci.
20(3): 12.

 

Kunisaki, J. T. and Y. Sagawa. 1971. Intermittent mist
to root anthurium cuttings. Haw. Farm Sci. 20(3): 4-5.

Nakashone, H. Y. and H. Kamemoto. 1962. Anthurium
culture with emphasis on the effects of some induced
environments on growth and flowering. Haw. Agri. Expt.
Sta. Tech. Bul. No. 50.

Neal, Marie C. 1948. In gardens of Hawaii. Bernice P.
Bishop Museum Special Publication No. 40. Honolulu,
Hawaii.

28.

29.

30.

31.

32.

33.

34.

8

Poole, R. T. and D. B. McConnell. 1971. Effects of
shade levels and fertilization on flowering of
Anthurium andreanum "Nitta" and "Kaumana". Proc. Amer.
Soc. Hort. Sci. Trop. Region 15: 189-195.

 

, R. Sakuoka and J. A. Silva. 1968. Nutrition
of Anthurium andreanum. Proc. Amer. Soc. Hort. Sci.
Trop. Region. 12: 2844287

Raabe, R. D. 1966. Control of anthracnose on the
island of Hawaii. Haw. Farm Sci. 15(2): 3.

Rasmussen, H. P. and B. D. Knezek. 1971. Electron
microprobe: Techniques and uses in soil and plant
analysis. Mich. Agri. Expt. Sta. Journal Art. No. 5214.
East Lansing, Michigan.

Shirakawa, T. and D. P. watson. 1964. Improved methods
of handling anthurium flowers. Haw. Farm Sci.
13(2): 1-3.

and R. R. Dedolph and D. P. Watson. 1965.
N-6 benzyladenine effects on chilling injury, respira-
tion and keeping quality of Anthurium andreanum. Proc.
Amer. Soc. Hort. Sci. 85: 642-646.

 

watson, D. P. and T. Shirakawa. 1966. Gross morphology
related to shelf life of anthurium flowers. Haw. Farm
Sci. 15(2): 2-3.

 

 

 

AN

SECTION I
ANATOMICAL STUDY OF THE ANTHURIUM PLANT,

ANTHURIUM ANDREANUM, L.

 

Section I: Anatomical Study of the
Anthurium Plant,
Anthurium andreanum, L.

 

Literature Review

Christensen (2) has shown that Anthurium scherzerianum,

 

 

and Anthurium andreanum, Lind., have a juvenile phase

 

followed by a generative phase. The difference can be seen
at the base of the petiole. Their observations were made

by dissection or removal of the petioles or dissection of
the shoot tips of 18 months to 2 year old seedling plants.
The juvenile stage formed leaves with a short sheath and a
vegetative bud at the axil; while the generative stage had a
flower bud at the axil and no leaf sheath. Instead the
stipules, which make up the sheath, cover the axil and the
upper part of the petiole base and protect the flower bud.

Watson and Shirakawa (9) examined flowers of Anthurium

 

andreanum, L., cv. Ozaki Red, at four stages of maturity

 

to observe the gross morphology and arrangement of indi-
vidual flowers. The study related the stages of develop-
ment of the flowers to stages of maturity and its relation-
ship to water loss and vase life. Greatest water loss
occurred when the stigma was receptive, open and thus

most vulnerable to water loss. Flowers, where the stigma

was either enclosed within the tepals or the stigma no

10

11

longer receptive and had become dehydrated, were less
vulnerable to loss of water from the spadix and, therefore,
had better vase life. Spadixes dipped in paraffin at 70°C
to prevent water loss were consistently turgid and fresher
after 12 days than untreated flowers.

Sharma and Bhattacharyya (8) found the chromosome
number of Anthurium andreanum, Lind., to be 2N=30.

Kamemoto and Nakasone (6) investigated the genetics

of flowers in Anthurium andreanum, Lind. and they found

 

Red, Rr, to be dominant to orange, R0; white, rr, breeds
true to white. A red in heterozygous condition, RrRO,
crossed to a white, rr, will give red, Rrr and coral pink,
R°r, in equal porportions. Orange, R°R°, crossed to white,
rr, results in only coral pink, R°r. Red or nonred spadix
color is simply inherited and interacts to some extent with
spathe color. Development of chlorophyll in the spathe is
simply inherited, but the transmission of sucker productiv-
ity and doubleness of spathe is not clearly defined.
Akamine and Goo (1) found a correlation between
physical characteristic and vase life of anthurium flowers.
They posited that the physical characteristics of petiole
length, diameter and flower weight of anthuriums were
significantly correlated with vase life, that is, the
greater the magnitude of the parameters, the shorter the

vase life.

12

Materials and Methods

Whole plant observation coupled with dissection, light

microscope and scanning electron microscope observations

were used to characterize the anthurium both morpholog-

ically and anatomically. Cv. Ozaki Red was used in the

investigation.

1.

Dissection of whole plants: Fresh matured
anthurium plants were dissected systematically
from root to stem, leaf and flower. The gross
anatomy was studied and recorded. A Bausch-
Lomb Stereo Zoom dissecting microscope and a
Hasting Triplett 14X hand lens (Bausch and

Lomb Optical Co., Rochester, N. Y.) aided in

the study. Selected photographs were taken of
the various plant parts with the aid of the
dissecting microscope.

Light microscope: Microscopic histological
procedures as outlined by Jensen (5) were followed
to prepare permanent slides of anthurium tissues.
Fresh anthurium tissues, usually 5mm2, taken

from various parts of the plant were killed and
fixed in a formalin-acetic acid-alcohol (FAA)
solution by placing them under partial vacuum for
one hour to ensure thorough penetration of the
FAA. FAA was prepared by mixing the following

solutions:

13
90 m1 of 50% ethyl alcohol
5 ml of glacial acetic acid
5 m1 of commercial formalin (40%)
The solution was filtered once with Whatman #1

filter paper.

The tissues were left in the FAA for a minimum
of 24 hours and then dehydrated through the
standard tert-butyl alcohol series. The series

consisted of the following solutions:

 

 

 

Solution Alcohol Content Distilled water
ETOH TBA
Number (ml) (m1) fl) (ml)
1 50 40 10 50
2 70 50 20 30
3 85 50 35 15
4 95 45 55 --
5 100 25 75 --
5 TBA -- 100 --

The tissues were left in solutions #1 to #5 for a
minimum of 4 hours and in the 100% TBA for a minimum of 12
hours, where it was repeated 3 times.

The tissues were then infiltrated with Tissuemat using
the following procedure:

A. Tissue placed in 5 mm x 20 mm vials in TBA.

Tissues were completely covered by the solution.
B. The content of the TBA was marked with a thin
strip of masking tape on the outside of the
vial. This was done to monitor the exact level
of the TBA as Tissuemat is slowly melted into

the solution.

14

C. Whatman #1 filter paper, cut and shaped like a
cone was suspended near the open end of each
vial (Figure 1). Tiny pieces of Tissuemat
were placed in the filter paper; the vial capped
securely with a rubber stopper and the vials
were placed in an oven at 60°F. The melted wax
dripped through the filter paper, into the TBA
and slowly infiltrated the tissue.

D. The Tissuemat was refilled at two hour intervals
until the content of melted Tissuemat and TBA
doubled (using the marking placed earlier on the
outside of the vial as a guide) or until enough
Tissuemat was melted to assure the complete
coverage of the tissues when the TBA was evap-
orated. This process took approximately 16 hours.

E. Once assured of the Tissuemat contentzhithe vial,
the stopper was removed and the TBA evaporated in
the oven at 60°F for 12 hours or until no TBA

could be detected.

The tissue was embedded in paper embedding boats and
mounted on wooden blocks 15mm x 15mm x 25mm, sectioned at
8 micrometers with a Leitz Wetzlar rotary microtome and
affixed to glass slides with Haupt's adhesive. The sections
were spread with 4% formalin on a slide warmer at 46°C.
The sections were positioned on the slide with a needle and
the solution removed withzapaper towel. The slides were

left overnight on the slide warmer and subsequently stained

15

Figure 1. An apparatus used for Tissuemat
infiltration.

 

16

RUBBER STOPPER

FILTER PAPER

TISSUEMAT PARAFFIN

DROP OF NELTED TISSUENAT

  

MARKER FOR TBA LEVEL
/
PLANT TISSUE

 

TBA

 

 

with safr;
staining 1

A.

 

17

with safranin and counterstained with fast green. The

staining procedure was as follows:

A.

Removed the paraffin from the sections by placing
the slideszhixylene for five minutes and then in
a 1:1 mixture of xylene and absolute alcohol for
an additional five minutes.

Partially rehydrated the sections by passing them
through a series of alcohols of decreasing con-
centration: absolute, 95%, 70%, and 50% (five
minutes in each).

Stained in safranin for three hours (1% safranin
in 95% alcohol and dilute with an equal amount of
distilled water).

Washed in water, differentiated with acidified
70% alcohol and passed rapidly through 95% and
absolute alcohol.

Counterstained with fast green (0.5% solution in
50% clove oil, 50% alcohol) for 30 seconds.
Differentiated the fast green in a mixture of 50%
clove oil, 25% absolute alcohol and 25% xylene.
Two changes were made at 10 minutes each.

Placed them in xylene, making 3 changes of 15

minutes each.

Permanent mounts were made with 22 x 50 mm cover glass

and Lipshaw Cover Glass Mounting Media. The tissues were

studied with an Olympus compound microscope and pertinent

data recorded. Photomicrographs were taken through a Zeiss

photomicrosc0pe.

l8
3. Scanning electron microscope (SEM): Tissue
preparation for viewing with the SEM as outlined
by Rasmussen and Hooper (7) was followed. Fresh
tissues about 5 mm2 were killed and fixed in FAA.
They were dehydrated in a graded ethanol-water
series for 30 minutes as follows:
25% ethyl alcohol
50% ethyl alcohol
75% ethyl alcohol
95% ethyl alcohol
100% ethyl alcohol
The alcohol was then removed by an iso-amyl acetate
series of:
25% iso-amyl acetate
50% iso-amyl acetate
75% iso-amyl acetate
100% iso-amyl acetate 2 times
The tissues were critical point dried in a Denton
DCP-l using liquid C02 at 1,650 pounds per square inch for
10 minutes. The tissues were mounted on 15 mm round cover
glasses using a drop of Tube Koat (G. C. Electronics Company,
Rockford, Illinois). Tube Koat, a carbon compound, was
used as a glue to insure electrical conductivity. The
tissues were coated with approximately 20 nm carbon followed
by 20 to 40 nm of Au-Pd (60%-40%) by evaporation. The

coated tissues were studied in the SEM (Advanced Metal

 

Research

photomic

Gross mc

 

shown in
perennia
Shade wfl'

3 to 6 I

are 51111;
maturit;
to a th.
a Pulvi~
Stipule

Th
1 Cm ap
much as

and the

I
OZaki 1P

has lor
tion ar
produce
leaves I

A
099031.

 

the le

 

19
Research Model 900) at 21 kV accelerating potential. SEM

photomicrographs were also taken.

Results and Discussions

Gross morphology: A typical Anthurium andreanum plant is

 

 

shown in Figure 2. It is a relatively low growing
perennial-herbaceous plant that thrives best in 60-80%
shade with cool temperatures and high humidity. It has
3 to 6 leaves in clusters rising from the stem. The leaves
are simple, chordate with smooth lamina margins and at
maturity are usually 20 by 35 cm. The leaves are attached
to a thin, long cylindrical petiole which forms a sheath or
a pulvinus at its base which is fused around the stem.
Stipules are borne at the sheath junction on mature leaves.

The usual short, thick stem has nodes approximately
1 cm apart. However, it may be a long thin stem with as
much as 15 cm between nodes: this depends on the cultivar
and the environment they grow in. For example, cv. Red
Ozaki has short, stocky internodes, whereas, cv. Nitta Orange
has long, thin internodes. Increased shade promotes elonga-
tion and long internodes, while high light intensities
produce short internodes regardless of cultivar. The
leaves have a spiral arrangement on the stem.

A vegetative lateral bud is formed at alternate nodes,
opposite each leaf junction. The vegetative buds are not
found in the leaf axils, but are spirally arranged opposite

the leaves.

 

Figure 2.

20

A mature anthurium plant,
Anthruium andreanum, L.,
cv. Red Ozaki.

 

—_m— H _

 

—fi_——- *w— .-

an- _——__ —-————_ fly

21

 

        

 

(I

==m I‘mleld

.n.....................1‘1. us<um

 

 

 

 

5:3... / I “:23

 

53mm

22

The flower buds are borne at each leaf axil. Mature
flowers consist of a conspicuous bract (spathe) joined to
the base of a cylindrical spike or rachis (spadix) and
attached to a long pedicel. The botanical flowers are
minute, perfect and densely cover the spadix. There are
approximately 300 flowers spirally and closely arranged on
the spadix. The fruits are small, globose, yellow berries
closely arranged on the spadix. There are two seeds per
berry which germinate approximately two weeks after sowing.
The first flower is produced about 18 months after
germination.

The roots are adventitious, all originating from the
stem. They are cylindrical, thick and fleshy and emerge
from the stem at each node, either below the vegetative
bud, the leaf, or both. The primary roots branch to form
secondary and tertiary roots. Root hairs are present 40-
50 mm from the meristematic root apex.

Two scale leaves or budscales are formed at each node.
One between the flower pedicel and the stem, the other
covering the lateral vegetative bud. The former is the
remains of the budscale which originally covered the apical
meristem of the plant.

Plant propagation is by seed and more often vegetatively
by either suckers (lateral shoots), stem cuttings or ter-
minal cuttings.

Christensen (2) reported that Anthurium andreanum when

 

propagated by seed, goes through a juvenile phase, followed

23
by a generative phase. This study supports that finding.
The difference in phases can be seen at the base of the
leaf petiole. In seedlings the first leaves formed have
short petiole sheaths with a vegetative bud at the axil.
As the plant matures a flower bud forms at the axil of the
leaf with no petiole sheath. Instead, the stipules form
the sheath covering the axil and upper part of the petiole
base, thus protecting the flower bud. In addition, the
lateral vegetative bud deve10ps opposite the leaf
attachment.

Crystals were found throughout the plant. The
crystals were primarily the druse and raphide forms
(Figure 3A). The druse crystal as viewed with the SEM is
shown in Figure 33. Energy dispersive X-ray analysis
indicates it is high in Ca, probably as calcium oxylate.
These crystals in the anthurium is in accord with most
plants in the aroid family (4).

A most characteristic feature of the anthurium is the
thick layer of cuticle on the epidermal cells of the aerial
parts of the plant. The cuticle is of particular interest
in anthurium research due to the problems of penetrability
Iby chemicals applied as mineral nutrients, fungicides or
herbicides (Figure 4). It also has a positive nature of

protection .

.gnatomy of the flower: The "commercial flower" of

 

.Anthurium andreanum consists of three main parts: 1) the

 

jpedicel or the flower stem; 2) the bracteoles or the

fill.“ V‘sfl'V'U—Wm

Figure 3.

24

Scanning electron micrograph of
rhombohedral crystals in the
anthurium. A left=raphide,
right-druse (arrows), X30; B=SEM
photomicrograph of the druse,
X500. r=raphide crystal, d=druse
crystal.

' -- _A\‘_r

LAMA-At."

 

 

arm ._._._- - -_- : :m_a_-‘I‘II

Figure 4.

26

A scanning electron micrograph
of the upper cuticle of an
anthurium leaf. X2050.

 

 

 

 

 

 

 

 

.,_‘-. -_ -f’w-

27

..
.

4o

 

28
spathe: and 3) the spadix. All three parts are attached
at a common junction. An anthurium flower is shown in
Figure 5A. The botanical flower is perfect, small and
borne on the spadix on a central rachis (Figure 5B). The
proximal flowerscxithe spadix develop first progressing
toward the apex. The surface anatomy and t0p view of the
botanical flower is shown in Figures 5C and D. Four
fleshy perianth segments (tepals) envelop four stamens and
a fleshy pistal. Each stamen is located opposite a tepal
and pressed firmly against the pistal. A cross section of
a botanical flower and spadix is shown in Figures 5E and
F. At the peripheral margin of the spadix the tepals form
the floral cavity and are attached to the central rachis.
The floral cavity contains the stamen and carpels. Anatom-
ically the tepals have an outer epidermal cell layer over
ground parenchyma cells. Vascular bundles are interspersed
throughout the ground cells. Many tepal cells contain
druse and raphide crystals, more than any other tissues in
the anthurium plant. These crystals are found scattered
throughout the plant, even in some epidermal cells.

The stamen has a flat filament with a single vascular
bundle in the center, which traverse the entire filament
and ends blindly in the connective tissue located between
the two anthers. The anther is bilobed with each lobe
containing two locules (Figures 6A and B). The outermost
layer is a one celled epidermis. The subepidermal layer

is the endothecium and the innermost layer the tapetum

firmnww-nnd

Figure 5.

29

The anatomy and morphology of the anthurium
flower. A=anthurium flower with its spathe

and spadix, X.75; B=spadix with its botanical
flowers, X1; C=surface morphology of the
botanical flower, SEM photomicrograph, x30;
D=botanical flower, X10; E=cross section of the
botanical flower, photomicrograph, x30.

F=cross section of the spadix x10; t=tepal,
s=stigma, a=anther, pt=pista1, fc=floral cavityr

30

 

B botanical Flowers

 

 

‘ . ‘O: I.
ii—c'samsrsa

Figure 6.

31

Photomicograph of the stamen of anthurium.
A=longitudinal section of the stamen, X16;
E=cross section of the anther, X32; C=pollen
being shed on the spadix, X2; D=anther

with its pollen, X10. pg=pollen grain,
a=anther, f=fi1ament, e=epidermal layer,
en=endothecium, lc=locule cavity, vb=
vascular bundle.

 

32

 
   

‘3‘!“ '2']

'0)?“ :3; a

33
(not shown in the photograph), is a nutritive tissue
frequently composed of multinucleate cells. The tissue
between the endothecium and tapetum is often crushed during
development of the pollen sac. The developing sac is evi-
dent in the locules. Mature pollen can be detected as a
white powdery mass on the spadix (Figures 6C and D).

The anthurium pistal is shown in Figure 7. The ovary
is hypogynous, as the tepals and androecium arise from the
receptacle below the gynoecium (Figure 5E). It is a
syncarpous gynoecium (more than one carpel) with two
carpels fused together. Each carpel has a single locule
with a single ovule. Therefore, each berry has two ovules
or embryos (Figure 73). Within the ovary the wall and the
locule cavity can be identified (Figures 73 and C). Ovule
placentation occurs in the center of the ovary where the
carpellary margins meet (axile placentation). The single
sessile style is an upward prolongation of the carpel with
two stigmas that elongate into papillae, feather-like short
hairs (Figure 7D). The stigma has glandular secreting
tissue and the stigmatic fluid provides a suitable medium
for pollen germination (3).

As the flower develops, the tip of the cylindric,
subconic pistal protrudes through an opening at the tip
junction of the tepals and the expanded style exposes the
stigmatic surface (Figure 7E). The stigma is receptive to
pollination when a sticky, translucent, stigmatic fluid is
secreted at the tip of each floWer on the spadix (Figures

7F and G). Pollination can be accomplished by collection

 

Figure 7.

34

Photomicrograph of the gynoecium of anthurium.
A=pista1 (arrow), X5; B=1ongitudinal section
of the pistal, X50; C=longitudinal section of
the ovary, X125; D=1ongitudinal section of the
stigma, X150; E=top View of the stigmatic area,
X600; F=spadix with receptive stigma, X1;
G=receptive stigma with its stigmatic fluid,
X5; H=berry and two embryos, X1. o=ovu1e,
cw=carpel wall, lc=locu1e cavity, de=developing
embryo, ow=ovary wall, p=papi11ae hairs of the
stigma, sf=stigmatic fluid, s=stigma, b=berry,
em=embryo.

35

  
  

 

   

36
of pollen on a soft brush and applying it to a receptive
stigma on the spadix. Natural pollination also takes
place with insects.
Each fertilized ovary enlarges to form a single berry
and each berry contains one or two embryos (Figure 7H).
The seed develops and is ready for sowing in six months.

The Anthurium andreanum flower is protogynyous. After the

 

 

stigmas on the spadix protrude through the tepals they
become receptive, dehydrate and shrink. The anthers grow
around the pistal and shed their pollen above the tepals.
The time lapse between the receptive stigma and the pollen
maturity is about a week.

The anthurium spathe,usually 12 cm x 14cm, is a modi-
fied leaf or bract. It is normally chordate, but may assume
other shapes (Figure 8). They are usually simple, but some
cultivars have a compound spathe. The spathe, in various

colors, is the product for which the Anthurium andreanum is

 

grown commercially in Hawaii. The prominent veins of the
spathe are parallel and originate at the spadix-spathe-
pedicel junction and diverge following spathe shape. The
parallel veins are interconnected by smaller transverse
veins. The thickness of the spathe is approximately 400-
600 microns.

The upper and lower surfaces of the spathe are covered
with a heavy layer of cuticle and except for a slightly
lighter color of the lower epidermis, there is little
anatomical differences between surfaces. There are 6-7

stomates per square mm on the lower surface and contrary

F'-

 

a'nzm ulnar-nu”

Figure 8.

37

The various spathe shapes of the
anthurium flower. A=cv.Ozaki Red;
B=UH 17 (pink obake); C=UH 16
(pink obake); D=cv. Anuenue;

B=UH 100 (Red Double); F=cv. Red
Elf.

 

 

 

 

 

38

 

39
to the report by Watson and Shirakawa (9) a few stomates
are present on the upper surface (Figure 9A). The
stomates are common to the epidermal layer.

In cross section (Figure 98) just below the upper
cuticle layer of the spathe is a single cell layer of
epidermal cells. These cells are broad, rhombical and
usually five sided, unlike typical flat, rectangular
shaped epidermal cells of most plants. Beneath the epi-
dermis is a single or double layer of isodiametric
hypodermal cells. Similarly the lower epidermis is also
a single cell layer composed of more typically flat,
rectangular cells. Inside the lower epidermis is a single
layer of hypodermal cells. Between the upper and lower
hypodermis layers of cells are found 10-12 irregularly
arranged layers of spongy parenchyma cells. Vascular
bundles are dispersed at uniform intervals throughout the
spathe. Microscopic observation of fresh samples indicated
that the anthocyanin pigments of the spathe are concen-
trated in the hypodermis cells. Both the epidermal and
spongy parenchyma cells are devoid of anthocyanin.

The mature flower stem or pedicel is smooth, cylindri-
cal and approximately 40-60 cm in length and 0.5 cm in
diameter. The outer surface of the epidermis is covered
with a thick, waxy cuticle layer with some stomates
(Figure 10A).

In cross section the pedicel (Figures 108 and C) is

anatomically similar to a monocot stem. A single layer of

 

:1”

I

Figure 9.

40

Photomicrograph and SEM micrograph of anthurium.
A=stomata on the upper surface of the spathe
(arrow), X2000; E=cross-section of the spathe,
X16. ep=epidermis, hy=hypodermis, sp=spongy
parenchyma tissue, vb=vascular bundle.

 

 

41

 

Figure 9

*7

Figure 10.

42

Photomicrographs and SEM micorgraph of the
anthurium pedicle. A=stomata on the pedicel
surface (arrow), X1600; E=cross section of

the pedicel, X30; C=cross section of the
pedicel, X60; D=cross section of the vascular
bundle, X300. sl=sclerified parenchyma tissue,
vb=vascular bundle, cx=cortex, ep=epidermis,
ph=phloem, x=xylem, ss=sheath of sclerenchyma
cells. ‘

43

 

3,1,1"; 39 J
10.0... :6". n 'l :
.r"..'.°£ :.:G‘ 1.4,... 5"?3}
.33. 3-

   

44
epidermal cells form the outer surface subtended by the
cortex. A layer of subepidermal sclerified parenchyma
cells separate the cortex and the ground parenchyma cells.
The vascular bundles, consisting of the phloem and xylem
are dispersed throughout the stem among the ground paren-
chyma cells (Figure 10D). These vascular bundles are
enclosed in sheaths of sclerenchyma tissues. The pith is

continuous and composed of ground parenchyma cells.

Anatomy of the leaf: The leaf blade is simple, green,

 

chordate with smooth margins and is approximately 20 cm
wide, 35 cm long and 400-500 mm thick. Leaf venation is
parallel with the main veins emerging from the base of the
leaf at the petiole-blade junction. The veins extend out
to the leaf margins, converging or fusing at the leaf apex.
The major veins are interconnected by smaller veins in a
complex pattern. Stomatal pores are found only on the lower
leaf surface (Figure 11A). They are located common to the
epidermal cells with approximately 30-40 stomates per
square mm with an apparently random distribution. No
stipules, hairs or other appendages are evident on the leaf
blade or the petiole.

The blade consists of an upper and lower one-celled
epidermal layer (Figure 118). The upper epidermal cells
are thicker and isodiametric hishape, while the lower
epidermal cells are much flatter. Between the epidermal
layers is the mesophyll made up of one or two layers of

compact palisade parenchyma cells. The palisade cells are

 

 

 

 

1.1.”33.

 

 

Figure 11.

45

SEM and light photomicrograph of the leaf of
the anthurium. A=lower leaf surface with
stomata (arrows), X500; E=cross-section of

the leaf, X35; C=petiole with stomata (arrow),
X500; SEM photograph, X500; D=cross—section
of petiole, X50; E=cross-section of vascular
bundle, X60; F=logitudina1 section of petiole,
X200; sp=spongy parenchyma tissue, pp=palisade
parenchyma tissue, vb=vascular bundle, ep=
epidermis, sl=sclerified parenchyma cells,
cx=cortex, ph=phloem, x=xylem, ss=sheath of
sclerenchyma, cp=cortical parenchyma tissue.

46

 

   

.12..-. ..MN ..

:2} . 3.... 11.5.4...- :oo.. in}.
. .v ; IP11... H.110M’X
iriiDi-I

w: .‘\.
\w ~

It
5.111%. .‘PI...I

  

 

    

  
 

 
 

    

 

47
found only beneath the upper epidermal layer of the leaf
blade. The leaves are, therefore, dorsiventral or bifacial
resulting in a dark green appearance on the upper surface
and lighter green on the lower surface.

Vascular bundles of various sizes with adaxially
located xylem and thick sclerenchyma fibers are dispersed
throughout the mesophyll. Chloroplasts are present in all
mesophyll cells, but are concentrated in the palisade
cells.

The leaf petiole is cylindrical, smooth and usually
25-35 cm long. The base of the petiole forms a sheath
around the stem. Stipules at the end of the sheath protect
the flower bud in its early stages. The epidermal surface
is smooth and covered with a thick waxy cuticle. A few
stomates are dispersed irregularly on stem surface (Figure
11C).

A cross section of the petiole is shown in Figures
110 and E. The anatomical structure of the petiole is
similar to the pedicel described earlier. The epidermis
is composed of a single layer of cells and subtending it
is the cortex consisting of several layers of parenchyma
cells. A layer of subepidermal sclerified parenchyma cells
separate the cortex and the ground parenchyma cells.
Vascular bundles are dispersed throughout the ground
parenchyma tissue and are surrounded by a sclerenchymous
sheath. The pith is continuous and composed of isodia-

metric parenchyma cells.

48
Anatomy of the root: The roots are adventitious arising
from the stem. Each root emerges just below an axillary
bud or leaf at each node. Two roots normally emerge at
each node. There is no main tap root, but roots branch
readily into secondary, tertiary, etc., roots. The spongy,
fleshy roots are cylindrical, 0.5-1.0 mm in diameter. They
are epiphytic possessing some specialized contractile
roots with the capability of attaching themselves to the
surfaces, such as pots, tree bark, etc. Root hairs are
apparent about 40-501mnbehind the root cap in the region
of maturation (Figures 12A, B and C).

A longitudinal view of the root is shown in Figures
12D and E and as illustrated the outermost layer of cells
form the root cap. The root cap protects the meristematic
region located just behind the root cap. The cells are
thin walled, cubical in shape with large nuclei and dense
cytoplasm. Immediately adjacent is the region of elonga-
tion with no cell division, but where differentiation
begins and maturation continues. These cells have large
vacuoles, less dense cytoplasm and slightly thicker walls.
In the root hair zone the cells are more differentiated and
root hairs emerge from epidermal cells.

The root in cross section (Figure 12F) shows multiple
layers of epidermal cells. The multiseriate tissue
(multiple epidermis) called velamen is commonly found in
the Araceae family. The tissue consists of compact non-

living cells, often with secondary wall thickening. The

Figure 12.

49

Photomicrograph and SEM micrograph of the
anthurium root. A=root tip, X4; B=root cap,
X100; C=root hairs, X100; D=longitudinal
section of the root tip, X10; F=cross-section
of root tip, X60. rc=root cap, m=meristematic
region, el=region of elongation, mz=maturation
zone, x=xylem, p=phloem, ed=endodermis, cx=
cortex, ep=epidermis-

 

 

 

50

root cop/

 

root hcin/

 

 

 

 

 

 

51
function of the velamen is absorption, but studies by Dycus
and Knudson (3) on orchids indicated that its principle role
is mechanical protection and water conservation. The
cortex is located adjacent to the velamenvfixflllayers of
large, thin walled parenchyma cells. The innermost
cortical layer has smaller cells, which form the endo-
dermis. The xylem, phloem and parenchyma cells compose the

stele (Figure 12F).

Anatomy of the stem: The primary stem (Figure 13) is

 

cylindrical, fibrous and at maturity is approximately 2 cm
in diameter. Leaves are arranged spirally with a phyllo-
taxis quotient of 2/7 and a divergence angle of 102.9°.
The spiral pattern may develop either clockwise or counter
clockwise. Factors affecting this pattern is unknown.

The nodes appear as pairs as the internodes alternate
between "short" and "long" internodes. One node gives
rise to the lateral vegetative bud, while the other pro-
duces the leaf pediole. The short internode is about 0.5
cm long, while the long internode varies from 1 to 5 cm.
The long internode responds to environmental conditions
and its response varies with variety. In common usage,
the short internode is referred to as the "node" and the
long internode as the "internode." However, anatomically
the stem is structured as previously discussed. Stem
growth on a mature plant is about 5-10 cm per year, but

as it is a perennial, stems up to 4 m long are common. A

vegetative apical meristem develops at each lateral bud

 

Figure 13.

52

Diagram of the vegetative stem of the
anthurium.

53

I’JI

—- LONG INTERNODE

  
  
 
  
 

FLOWER BUD IN
LEAF SHEATH

 

 

n —— lATERAL BUD
.1. \ suont mrrnuoor

'~— LONG INTERNDDE

 

FLOWER PEDICEL

 

 

 

LEAF PETIDLE / LATERAL BUD

 

SHORT INTERNDDE ..
; NUDES
SCALE lEAF SCAR /

STEM

Figure 13

54
and stem terminal. This results in a network of stem
branching as the lateral shoots develop.

The anatomy of the stem is similar to the pedicel and
petiole discussed earlier. The stem has one layer of cells
forming the epidermis. The epidermis surrounds a cortex
layer but is much thicker in width than the pedicle or
petiole. A layer of sclerified parenchyma cells separate
the cortex from the ground parenchyma cells. Vascular
bundles enclosed in sheaths of sclerenchyma tissues are
dispersed in the ground tissues. The pith is continuous
and consists of thick walled parenchyma cells.

The vegetative apical meristem (Figure 14A) is off-
set from the terminal apex. This off-centered position is
caused by the spiral characteristic of the stem in which the
apical meristem is displaced terminally by the developing
vegetative primordias. The primordias are shown at the
periphery of the apex, possessing undifferentiated embryonic
cells.

At the meristematic apex are two layers of tunica cells
(Figure 143), which divide perpendicular to the surface of
the meristem (anticlinal division). Directly beneath the
tunica is the corpus where cell division occurs in multiple
planes (anticlinal and periclinal). The corpus adds bulk
to the apical portion of the shoot by increasing in volune,
whereas tunica maintains its continuity over the enlarging
body by surface growth.

To summarize the section: a study of the morphology and

anatomy of Anthurium andreanum, L., was conducted. It is a

 

 

I ‘1'! “U“

Figure 14.

55

Photomicrograph of the vegetative meristem of
the anthurium. A=longitudinal section of the
meristem (arrow), X15; B=en1arged view of the
meristematic region (arrow), X60. lp=leaf
primordia, fp=flower primordia, ls=leaf scale
primordia, la=lateral shoot primordia, tu=
tunica, co=corpus.

 

—

 

56

 

57
perennial-herbaceous monocotyledon in the family Araceae,
therefore, possesses typical monocot anatomical structures.
Of particular significance among its characteristics are:
1) possesses a juvenile phase as seedlings and a generative
phase at maturity. The juvenile phase has a lateral shoot
at the leaf axil, whereas, the generative phase has a
flower. The lateral shoot locates itself opposite the leaf
attachment; 2) the "commercial anthurium flower" has chor-
date spathe and a protruding spadix, whereas, the botanical
flowers are minute, perfect and borne on the spadix; 3) the
roots are covered with a multiple layer of epidermal cells
called velamen; 4) raphide and druse crystals are dispersed
throughout the entire plant tissue; and 5) the above ground
parts of the plant is covered with a thick layer of waxy

cuticle.

LITERATURE CITED

58

LITERATURE CITED

Akamine, E. K. and T. Goo. 1972. Relationships
between physical characteristics and vase life of
anthuriums. Haw. Farm Sci. 21(2): 9-10.

Christensen, V. O. 1971. Morphological studies on
the growth and flower formation of Anthurium
scherzerianum, Schott and Anthurium gndreanum, Lind.
Statens Forsogsvirksomhed i Tidssdr Planteval

75(6): 793-798.

 

 

Dycus, A. M. and L. Knudson. 1957. The role of the

velamen of the aerial roots of orchids. Bot. Gaz.
119:78-87.

Esau, Katherine. 1961. Anatomy of seed plants.
John Wiley and Sons, Inc., N.Y.

Jensen, W. A. 1962. Botanical histochemistry.
W. H. Freeman and Company, San Francisco, California.

Kamemoto, H. and H. Y. Nakasone. 1955. Improving
anthuriums through breeding. Haw. Farm Sci. 3(3): 4-5.

Rasmussen, H. P. and G. R. Hooper. 1974. Electron
Optics: Principles, techniques and application in
horticulture. Hort. Science 9(5): 425-433.

Sharmg, A. K. and U. C. Bhattachryya. 1961. Structure
and behavior of chromosomes in species of anthurium
with special reference to the accessory chromosomes.
Proc. Bio. Sci. 27:317-328.

Watson, D. P. and T. Shirakawa. 1966. Gross

morphology related to shelf life of anthurium flowers.
Haw. Farm Sci. 15(2): 2-3.

59

SECTION II
COLOR BREAKDOWN IN ANTHURIUM FLOWERS,

ANTHURIUM ANDREANUM, L.

 

60

Section II: Color Breakdown in Anthurium Flowers,
Anthurium andreanum, L.

 

Literature Review

In 1972 the Hawaii Plant Disease Clinic, College of
Tropical Agriculture identified the color breakdown or
water-soaked disorder of the anthurium spathe as oedema.

It was believed to be caused by excessive rains which
hindered normal transpiration of the plant, thereby causing
cellular injury (Figure l).

Oedema or intumescence is a blister-like, abnormal
protuberant out-growth on plants, occurring more frequently
on leaves than on other plant parts. The abnormality may
occur on stems, but rarely on the botanical flowers or
fruits (45). Atkinson in 1893 (1) reported oedema on

tomatoes. Subsequently oedema was reported on Hibiscus

 

vitifoluis, L. (5), on potatoes (6), manihot (45), poplar

 

(17, 21, and 22), grape (44), tobacco (11), geraniums (3),
and eggplants (7).

Anatomically, oedema has been shown to be an abnormal-
ity due to hypertrophy of the palisade cells and spongy
parenchyma (5, 6, and 45). This caused rupturing of the
epidermal cell layerscnithe leaf or stem surface resulting

in oedema or intumescence symptoms. In Pelargonium

 

hortorum, Ait., hypertrophied cells were confined to the

 

61

v
m

Figure l.

62

Anthurium flowers showing various stages of
the color breakdown disorder. A=norma1 flower;
B=flower with lobe section showing water-
soaked lesions; C=flower with lobe section
showing the lesions coalesced to effect the
entire lobe area; D=flower with lobe section
turned brown and necrotic. Cv. Ozaki Red.

 

 

 

 

63

 

64
spongy parenchyma cells, filling the substomatal cavity
and oedematic structures appeared only on the lower epi-
dermis of the older leaves (3).

In all the reports cited, the abnormality was
attributed to physiological rather than pathological con-
ditions. It was generally concluded that hypertrophy of
the palisade and spongy parenchyma cells resulted when
water absorption exceeded transpiration, causing water to
accumulate in the tissues. A continuous water supply to
a warm, moist soil with cool night temperatures predis-
posed plants to oedema (3). Other environmental factors,
e.g., light intensity, light quality and mineral concen-
tration in the plant were studied, however, plant-water
relationship was found to be the main factor causing
oedema (27, 28, and 29).

According to Chupp and Sherf (4), oedema may be
caused by any agent which stimulates groups of inner cells
to grow abnormally. This condition was accentuated when
moisture was high, soil temperature was high and air
temperature was low. These conditions occur on a cool
night after several warm rainy days when relative humidity
is near 100%. Under these conditions the roots absorb
water at a rapid rate exceeding that lost through trans-
piration. The result was an oversaturation of the cells
with water, creating enough pressure to rupture the
epidermis.

Similar physiological diseases due to moisture

relationship have been reported: blossom-end rot and black

65

seed of tomatoes (8, 41, and 43), withertop of flax (18),
bitter-pit of apples (38), blasted buds of lilies (37),
crinkle leaf of cotton (31), internal browning of brussel
sprouts (24), brownheart of escarole (25), internal
tipburn of cabbage (26), tipburn of lettuce (42), brown
rib of lettuce (15), blackheart of celery (10), and
blossom-end rot of guava (39). Each disorder cited above
was corrected or prevented by the application of Ca.

Sprague (40) has reported that Ca in plant cell walls
forms a protective ”sieve" for nutrient to seep through in
passing into cells. It also acts as a cement (calcium
pectate) between cell walls. Stackman (36) reported that
Ca deficiency was invariably associated with the break-
down of cell walls. Rasmussen (33) reported that Ca
deficiency weakened tobacco leaves, causing the constituent
cells to separate under little stress. Microscopic studies
showed that tobacco leaf tearing in tender leaf occurred
in an irregular pattern between cells. When Ca was present
in sufficient quantity, tearing occurred through cells.
However, the total Ca content was approximately the same in
normal and tender leaves. Further studies suggested that
there may be equal quantities of Ca in the sum of
pectate, crystal and soluable Ca in both normal and tender
leaf tobacco, but there is less in the pectate of the cell
walls of the tender leaf.

Lange (19) using beans, found that the lack of aera-

tion for 12 hr caused as much as 49% decrease in Ca uptake,

66
but increased the P uptake as much as five times. Lange
also found Ca uptake highest at pH 7.0. Ca accumulation by
apple stems and leaves (46) was less at 5°C than at 20°C,
30°C and 40°C. The maximum Ca translocation was between
15°C to 30°C. In peas (13) decreasing the temperature
from 20°C to 0°C during a 24 hr period decreased both water
and Ca uptake. These studies indicate that aeration, pH
and temperature affect Ca uptake.

Crowns of healthy strawberry plants (20) had a lower
percentage of ash and a higher percentage of Ca than other
parts of the plant. The roots of diseased plants had low
Ca levels, while the leaf interior had higher Ca levels than
the serrated edges. On the other hand, the serrated edges
of the leaves of healthy plants were higher in Ca than the
rest of the leaf. This study indicated that Ca content does
vary in different parts of the plant and even within the
same leaf. It may also vary between diseased and healthy
plants.

Moser (30) growing soybeans and sorghum in sand culture
showed that Ca concentration at the various pH levels
(pH 3.8-4.2, 5.0-5.2, and 6.0-6.5) was a more important
factor on Ca uptake than pH levels. His results showed
that increased supply of Ca influenced directly the concen-
tration of Ca within the crop irrespective of pH.
Lundegardh (23) found available Ca reduced from 25 mgms
to 12.5 mgms per liter when the pH of the soil was changed
from pH 7.0 to 5.0. He attributed this effect to chemical

precipitation of phosphate and other minor elements with Ca.

67
Materials and Methods

The problem of the color-breakdown in the spathe of
the anthurium flower was approached using the following
methods:

1. Isolation and inoculation of microorganisms from

diseased tissue.
2. Chemical elemental analysis of tissue from

affected and normal healthy flowers.

Isolation and inoculation of microorganism from diseased
tissue:

One 5 mm2 tissue section was taken from the water-
soaked and color-breakdown area of ten anthurium spathes,
cv. Ozaki Red. The tissue sections were surface sterili-
zed with 0.8% sodium hypochloride for 3 minutes, rinsed 3
times with sterile distilled water and cultured on potato
dextrose agar plates.

A fungus, identified as Colletochrichum gloeosporioides
was isolated from the tissues. Spore suspensions of asci
and conidia stages were prepared with distilled water and
sprayed with a hand atomizer on the anthurium spathes of
the cv. Ozaki Red. Treatments were as follows: 1. Ten
flowers sprayed with asci suspension; 2. Ten flowers
sprayed with conidia suspension; and 3. Ten flowers
sprayed with distilled water. The flowers were placed in
the greenhouse under a continuous fine mist for 48 hr in
the shade at 80°F. Daily observations were made to deter-

mine development of incipient water-soaked or color-

breakdown lesions.

68
Chemical elemental analysis of anthurium tissues from
affected and normal flowers:

Eighty, 75 percent mature flowers and 80 associated
leaves on the same axil of the stem, were collected from
the anthurium cv. Ozaki Red. Seventy-five percent mature
flowers were distinguished by the degree of color change
of the spadix, e.g., when the lower 3/4 of the spadix had
changed from reddish-orange to light pink in cv. Ozaki
Red. Forty of the flowers showed symptoms of the spathe
color-breakdown, while forty appeared to be normal healthy
flowers. The flowers and leaves were divided into four
treatments: 1. Color-breakdown flowers; 2. Leaves of
color-breakdown flowers; 3. Normal flowers; and 4. Leaves
of normal flowers. The 40 flowers and 40 leaves making up
each treatment were further divided into 4 samples of 10
flowers and 10 leaves (4 reps).

The samples were rinsed with distilled water and air-
dried. The spadix and pedicel were discarded and only the
spathe analyzed; similarly, only the leaf blade was
analyzed. Fresh and dry weights were taken and percent
moisture determined. The samples were dried in a force air
oven at 75°C for 48 hr, after which they were ground to a
fine powder in a Spex Industries Ball Mill.

Levels for N, P, K, Ca, Mg, B, and SiOz were deter-
mined as follows:

Nitrogen: the standard Kjeldahl total nitrogen method

 

as outlined by Horwitz (12).

69

Phosphorus, potassium, calcium and magnesium: 5 grams

 

of the ground sample were ashed overnight in a muffle
furnace at 500°C. The ash was "wet" down with distilled
water and dissolved with 5 ml concentrated hydrochloric
acid and left standing for one hr. The solution was trans-
ferred to a 50 ml volumetric flask, made up to volume with
distilled water and left standing overnight. An aliquot
was diluted 10 times and P was determined by the
Molybdophosphoric blue color method (14) (Klett-Summerson
Photoelectric Colorimeter, model 900.3). A second aliquot
from the 50 ml solution was diluted 10 times with 0.5%
Lanthanum solution and Ca and Mg were determined spectro-
photometrically (Perkin-Elmer Atomic Absorption
Spectrophotometer, Model 303). (The Lanthanum solution was
made by wetting down 58.65 gm of La203 with distilled water
and adding 250 m1 of concentrated HCl very slowly until the
material was dissolved. The solution was diluted to 1000
ml with distilled water to give a 5% lanthanum solution in
25% (v/v) HCl. Aliquot was taken from this stock solution
and diluted to a 0.5% Lanthanum solution. Lanthanum was
added to prevent interference from P and A1.) A third
aliquot from the 50 ml solution was diluted 10 times with
distilled water and analyzed for K by flame photometry
(Beckman DU Spectrophotometer).

Egrgn: Boron in ppm was determined by the Quinaliz-
arin method (12). Three ml of saturated calcium hydroxide

was added to 2 gm of the ground samples and left standing

70
for one hr. Samples were ashed overnight at 550°C. Ten
ml of 0.36N H2804 were added to the samples and left
standing for 2 hr. The solution was filtered with White
Ribbon Filter No. 589 into a 40 ml soft glass test tube
(non-pyrex) and shaken well. Two ml of the solution were
pipetted into an absorption test tube, ten ml quinalizarin
dye added (45 mg quinalizarin dye per liter of H2804),
mixed thoroughly, cooled to room temperature and measured
colorimetrically (Evelyn Colorimeter) at 595-660 micron.

Silicon dioxide: SiOz was determined by ashing 1 gm

 

of ground sample at 500°C overnight, dissolving the ash in
5 ml concentrated hydrochloric acid and drying over low heat
for three hr. The ash was again dissolved in 10 ml of 3N
hydrochloric acid and filtered with White Ribbon Filter

No. 589. The filter paper was ashed in a porcelain cruc-
ible for 3 hr at 700°C. The ash and crucible were cooled
in a dessicator over calcium chloride and weighed. Subse-
quently, the crucible was cleaned with a stiff brush to
remove the ash, re-ashed at 700°C for one hr, cooled in

the dessicator and reweighed to get the weight of the ash.
The ash weight was calculated as percent of original matter
and reported as percent SiOz. A blank sample on the filter
paper was similarly analyzed serving as a control for

filter paper impurities.

After preliminary experiments were carried out and a
hypothesis formulated, the following procedures were used

to verify the hypothesis:

71

1. Electron microprobe X-ray analysis of elements
in the tissues

2. Solution culture to induce the disorder symptoms

3. RadioisotOpes to study element distribution and
localization

4. Field test to verify the results

Electron micrgprobe X-rgy analysis:

 

Fresh samples, about 5 x 15 mm from an anthurium
spathe showing symptoms of color-breakdown and from normal,
healthy flowers were taken from cv. Ozaki Red for micro-
probe analysis. The tissues were taken from the lobe and
tip sections of the spathe for analysis, since the symptoms
of color-breakdown are localized1h1the lobe section of the
spathe (Figure 2). The cyrostat tissue preparation method
was used (34). The fresh tissue was immediately mounted
and frozen in Optimum Cutting-Temperature Compound (OCT
—15 to -30°C, Fisher Scientific Company). Sections 16
micrometers thick were cut at -16°C on the Model CTD-
International-Harris-Cryostat and placed at room tempera-
ture on polished carbon disc 50 mm in diameter. The OCT
served as the mounting medium. The samples were air-dried
and placed in an electron microprobe X-ray analyzer
(Applied Research Laboratory, model EMX-SM), operating at
21 kV accelerating voltage and 0.056 micro amperes sample
current. No carbon conductive coating was used with
sections of this thickness. Calcium X-ray emission counts

as well as graphic line profile analysis of K(2000)\),

72

Figure 2. Illustration of sampling of tissues from lobe
and tip sections of the anthurium Spathe.

53...:

”225% a:

 

”228mm :3

74

P(200A), Na(200A) and Ca(2007\) were taken and recorded.

Solution culture:

 

This method was used only after previous experiments
indicated that calcium levels may cause color-breakdown in
the spathe. Of prime interest was inducement of the color-
breakdown disorder by eliminating calcium in the nutrient
solution and/or by varying the pH levels of a complete
nutrient solution to alter chemically the availability of
calcium for uptake by the plants.

Twenty-eight mature anthurium plants of the cv. Ozaki
Red were planted in coarse perlite media in 8" plastic
pots. The pots were seated into a completely enclosed
wooden box as illustrasted in Figure 3. The peat moss was
kept moist at all times providing 100% relative humidity
to the roots. A heating cable in the peat moss kept the
temperature at the root zone at 80°F. The purpose of the
box was to provide an environment of high humidity and
high root temperature to increase metabolism, e.g., increase
nutrient, water uptake and growth. The box also prevented
drying of the roots as perlite is a poor water holding
media. The plants were given a nutrient solution as
described by Rasmussen, et a1. (34) with modifications
(Table I). Two hundred m1 of each treatment solution were
added three times a week. No additional watering was
necessary. pH levels were 3, 4, 5, 6, and 7 and were tested

as well as concentrations of 0, 100, and 200 ppm. The solutions

' I
"we!“

Figure 3.

75

Planting system used to grow anthuriums in
nutrient culture.

76

$23 5.;
825:3 m8: 2:

SS: “55.. z.
325.. 2:52.22

55 238; 3385

.....u......._........_......._.._ _. _. .. ~ 5 .. .. .. .5 Ce

 

.03 2
:8 SEE:

77

Table I. Solution Culture Composition, Concentrations
and Conditions used in the Calcium Experiment

Nutrient Culture

 

 

 

Major elements g/liter
*Ca(NO ) ° 4H20 1.18
KNO3 3 2 0.51
KH2PO4 0.14
M9504 ° 7H20 0.49
Iron sequestrene 0.0115

Minor elements g/liter
H3BO3 0.6

MnC12 ' 4H20 0.4
ZnSO4 0.05

Treatments

1. Complete, pH 6

2. 1/2 calcium, pH 6

3. No calcium, pH 6

4. Complete, pH 3

5. Complete, pH 4

6. Complete, pH 5

7. Complete, pH 7

*In the 1/2 calcium and No calcium solutions, NH4NO
at 0.20 g/l and 0.40 g/l were added respectively to
compensate for the loss of N when Ca(NO3)2 - 4H20
was reduced or omitted.

78
were pre-mixed in 20 liter plastic carboys and pH readings
taken weekly. When necessary, the pH was adjusted With
15% H2804 and/or 15% NaOH. Weekly production data and
any observable nutritional deficiency symptoms were
recorded. Calcium content of the spathe and the leaf
associated with the flower was determined spectrOphoto-
metrically as described earlier. The spathe and leaf
blade were divided into lobe and tip sections and analyzed.

Radioisotopes:

 

Twelve mature anthurium plants cv. Ozaki Red, were
grown in three gallon plastic containers in aerated solu-
tion culture (Figure 4). Plants were grown in a fiberglass
greenhouse under 80% shade. Approximately 500 ft. candle
of fluorescent light was given nightly from 5 p.m. to
5 a.m. to increase plant metabolism.

Plants were grown in deionized water for 5 days and then
transferred to nutrient solution as given in Table I, with
the exception that Ca(NO3)2 - 4H20 was omitted and CaC12
added to make a solution at 5 X 10'4 molar. NH4NO3 was
added at 0.40 gm/l to compensate the loss of nitrogen by
omission of Ca(NO3)2 - 4H20. The solutions were adjusted
to pH 3, 5, 7, and 9. Three plants per treatment were grown
for 7 days at the end of which, the solution was replaced,
pH adjusted and 0.05 micro-curies per m1 of 45Ca added to
each solution. The plants were allowed to take=up 45Ca
for an additional 14 days. pH was checked twice a day and

adjusted for the duration of the experiment.

mi

'3.

 

 

 

. I
III'I'I‘V vl-r. . 14d.

 

7’-

Figure 4.

79

Diagram of apparatus used for radioisotope study.

80

 

23:: m
.3558 a: I

«E I

90000:.

 

 

 

we: 35o

 

 

 

 

 

as? Eq 8

:2: 2:52:26.

 

292:8 EwESz

59:3 85 288;

81

The plants were harvested at 14 days and prepared for
microautoradiography and SEM studies. Fresh tissue 5 x 15
mm was taken from the flower spathe and leaf blade associ-
ated with the flower at the same stem axil. Tissues of both
spathe and leaf were taken from the upper lobe area and from
the tip section.

The tissues were fixed in a 3:1 95% alcohol-acetic
acid solution (v/v) and carried through tert-butyl alcohol
dehydration and tissuemat embedding (16). Sections were
cut at 8 micrometers (American Optical Company, model 815
rotary microtome) and mounted on gelatin-chrome-alum coated
glass slides. The paraffin was removed with xylene and
microautoradiography procedures of Ficq (9) were followed.
The sections were coated with Kodak Nuclear Track NTBZ
emulsion and stored in the dark for proper exposure. After
the microautoradiographs were developed, fixed, washed,
air-dried and a cover glass mounted permanently with Pro-
Texx mounting media, the sections were viewed with phase
contrast or bright field microscopy to determine silver
localization in the tissue.

For SEM energy dispersive X-ray analysis, the paraffin
embedded tissue was cut at 10 micrometers, mounted on
gelatin-chrome alum coated polished aluminum disc (17 mm
in diameter) and the paraffin removed with xylene. The
sections were air-dried, coated with 20 nm of carbon by
evaporation and analyzed in a SEM (Cambridge Stereoscan S4

with Edax detector) (35).

82

Field test:

 

To determine the effect of calcium on the color-
breakdown disorder, a randomized block design with three
treatments and three replicates was used. Seventy—two
mature anthurium plants, cv. Ozaki Red, were planted in
6' x 12' plots in black cinder media under 75% Saran
shade. They were subjected to the following calcium treat-
ments: 1. Calcium nitrate at the rate of 200#/A/yr;

2. Calcium silicate at the rate of 1400#/A/yr; and 3. No
calcium. Total calcium was equally divided into 3 parts
by weight and given to the plants as a soil additive every
4 months.

Normal fertilizer program of Osmocote 14-14-14 was
given 3 times a year at the rate of 300 # N-P-K/A/yr and
standard pesticide practices were followed.

Data on normal, healthy flowers and flowers showing

color-breakdown symptoms on the spathe were recorded.

Results and Discussion
Anthurium flowers sprayed with asci and conidia sus-

pensions of Colletochrichum gloeogporioides showed no sign

 

of the color-breakdown disorder in any of the treatments.
The normal senescence symptoms, such as browning of the
spadix and eventual abscission of the flower from the stem
at the "stem-spathe-spadix junction" took place within 15
days after treatment. It was concluded that the fungus

Colletochrichum gloeogporioides, earlier isolated from the

 

color-breakdown spathe, was not the cause of that disorder

83
in anthuriums. Aragaki (2) writes:

Colletotrichum gloeosporioides is a common fungus
on tropical plants, especially on above ground
parts. It is recovered in very high frequency
from healthy-appearing tissues, so not surpris-
ingly, the rate of recovery from necrotic tissue
is even higher. Obviously, many of them are non—
pathogenic, and this has been verified by
innoculation tests. It is also a pathogenic
species, however, and will cause many important
diseases, including anthurium anthracnose. In
the years (I) spent on screening anthuriums for
anthracnose resistance, spathe lesions have been
few and far between, and not reproducible.
Approximately 50 isolates of g, gloeosporioides
(including several from spathe) have been
screened thus far.

 

Our results verify these results.

Elemental analysis of anthurium spathe and leaf
tissue from color-breakdown and normal flowers is given in
Table 2. Differences in Ca are evident. Color-breakdown
spathe and leaves have a lower Ca content (0.372% and 0.363%
reSpectively) than normal spathe and leaves (0.830% and
0.805% reSpectively). No other elements varied between
color-breakdown and normal plants. The data suggest that
low Ca and not oedema may cause the anthurium color-
breakdown disorder.

Electron microprobe X-ray analysis of a cross-section
of spathe tissue in both color breakdown and normal tissue
is given in Figure 5. A higher concentration of Ca is
evident in the normal flowers as compared to color-
breakdown flowers. Normal tissue had a higher concentra-
tion of Ca in the upper and lower epidermal tissue than in
intermediate tissues. Ca distribution in color-breakdown

tissue was uniformly low. Semiquantitative analysis for

84

Table 2. Elemental analysis of anthurium spathe and
leaf from color-breakdown and normal plants.*

 

B %
Spathe %N %P %K %Ca %Mg %SiO2 ppm moisture

 

Normal 1.55 .180 2.13 .830 .176 .10 16.0 13.17

 

Color-
break 1.76 .201 2.44 .372 .150 .12 14.0 13.78

 

Leaves

 

Normal 2.12 .178 2.01 .805 .266 .10 11.8 21.68

 

Color-
break 2.20 .173 1.98 .363 .196 .10 13.6 24.18

 

*Note: Each figure represents an average of 10 plants per
sample and analysis ran on 4 replicated samples.

Figure 5.

85

Microprobe line profile X-ray analysis of Ca in
the spathe of anthurium plants. A=color-break-
down flower, lobe section; B-color-breakdown
flower, tip section; C=normal flower, lobe
section; and D=normal flower, tip section. The
line scan proceeded from X, upper epidermis, to
Z, lower epidermis.

 

 

 

 

 

 

 

 

 

 

87
Ca in cross-sections of the upper, middle and lower sections
of the spathe tissue produced similar results (Table 3).
Lanning (20) found variation in Ca content of strawberry
within the same plant part and even in the same cross-
section. These data strengthen the hypothesis that low Ca
is the cause of the color-breakdown disorder in anthuriums.
Tissue exhibiting the color-breakdown had a lower Ca
content than normal tissue.

Nutrient culture studies carried out in Hawaii pro-
duced color—breakdown symptoms on the spathe after 6 months
of growth in nutrient solutions. Color-breakdown symptoms
occurred consistently in the "No Ca" treatment. Other
treatments such as 1/2 Ca and the various pH levels (pH
3, 4, 5, 6, and 7) showed no symptoms of color breakdown
after 12 months. However, the ”No Ca" treatment plants
which developed color breakdown symptoms produced normal
flowers when Ca was added. The disorder symptoms were as
follows: the spathe developed tiny water-soaked lesions or
discolored areas the size of pin headscnithe upper epi-
dermis on the lobe section of the spathe. These lesions
increased in number and size and eventually coalesced to
form a water-soaked mass or large color-breakdown lesion.
The lesions dehydrated, turned dark brown and died with an
upward roll of the necrotic tissue. In extreme cases, the
young leaves became chlorotic and were small and distorted.
The margins were irregular and frequently contained spotted

and necrotic areas. New leaves and flowers continue to

88

Table 3. Ca X-ray analysis of a cross-section of the
spathe of anthurium in normal and color-breakdown
flowers.

Ca distribution (counts/sec.)*

 

 

 

Normal flower Colorbreak flower
Upper epidermis area 922 76
Middle area 699 67
Lower epidermis area 679 124

*Note: Each figure represents an average of 9 counts
taken from 3 colorbreak and/or 3 normal flowers.

89

emerge, however, they die before they unfurl. The apical
meristem was obviously affected since lateral shoots began
to develop from the stem. The entire plant was stunted and
eventually died.

Elemental analysis of plants grown at various nutri-
ent and pH levels in the cultural solutions was made.
The plant was separated into lobe and tip sections of both
the spathe and leaf. The results are presented in Table 4.
Again there was less Ca in those plants exhibiting color-
breakdown (No Ca treatment), both in the spathe and in the
leaves as compared to all other treatments. The results
also indicate less Ca in the lobe than in the tip. This
further suggests that low Ca is the cause of the disorder.
The symptoms of this disorder are initiated and concen-
trated in the lobe section of the spathe. Calcium uptake
increased with increase in pH levels from 3 to 7.

Tissue samples of the lobe and tip of the spathe and
of plant leaves grown at various pH levels were analyzed
by energy dispersive X-ray analysis (Figure 6). The uptake
of Ca was lowest at pH 3, increased through pH 5, reached
a maximum at pH 7 and decreased again at pH 9. These
results are similar to other reports of the effect of pH
on Ca uptake (19 and 23). This pH data sheds some light on
the course of the color-breakdown disorder found in Hawaii's
anthurium nurseries. The major occurrence of the color-
breakdown disorder is in nurseries using volcanic black
cinder as a growing medium. The medium is a light, porous,

inert material which provides excellent drainage and

90

Table 4. Ca content of anthurium spathe and leaf
tissues grown at various nutrient andng levels.*

Ca concentration (%)

 

 

Flower Flower Leaf Leaf
Treatment lobe tip lobe tip
No Ca, pH 6** .060 .070 .130 .130
1/2 CA, pH 6 .320 .500 .830 .880
Complete, pH 3 .370 .570 .730 .790
Complete, pH 4 .430 .710 .850 .890
Complete, pH 5 .420 .700 .880 .880
Complete, pH 6 .520 .830 1.160 1.150
Complete, pH 7 .540 .850 1.200 1.215

*Note: Each figure is an average of 4 plants per sample.

**Note: Treatment with color-breakdown symptoms.

 

e
ILL.

Figure 6.

91

Energy dispersive X-ray analysis of anthurium
flower sections of plants grown at various
nutrient and pH levels. Photographs on the
left=1obe sections; right=tip sections.

A & B=pH 3; C & D=pH 5; E & F=pH 7; G & H=

pH 9.

 

 

 

 

_—_——M"—7

h_“—_‘——

92

 

 

 

 

 

 

 

 

93

aeration, is low-cost and readily available as it is a
by-product of the active Hawaiian volcanoes. Its wide
spread utilization was realized only after the development
of slow-release fertilizers. Generally, the ordinary
commercial inorganic fertilizers were not effective for
anthuriums because they were readily leached with the
heavy Hawaiian rains before the anthurium could utilize
the nutrients. Unfertilized, fresh volcanic black cinders
have a pH of 6.0 to 7.0. When used as a growth medium with
organic fertilizers the pH drops to near 4.0. The acid pH,
coupled with the fact that liming is not practiced in the
nurseries, may account for the color-breakdown spathe dis-
order problem in Hawaii. On the other hand, pH studies with
the nutrient solutions did not produce the color-breakdown
disorder (page 87). The minimum Ca requirement was not
established and even at pH 3.0 this may have been met in
a nutrient system. Each solution treatment contained
Ca(NO3)2-4H20 at 5 x 10"3 molar. Similar results were
reported by Noro (32) who found that olive trees fed "high"
levels of Ca showed no deficiency symptoms even at pH
4.0-4.5 and Moser (30) concluded that the Ca concentration
at various pH levels for soybeans and sorghum was a more
important factor than pH levels on uptake of Ca.

Microautoradiography with Ca45 showed that Ca was
primarily deposited and concentrated in the cell wall.

The practical aspect of any scientific work is of

utmost importance. Therefore, the hypothesis that Ca

94

deficiency was the cause of the disorder was tested in the
field. The field test results (Table 5) confirm that the
application of Ca will significantly reduce the incidence
of the color-breakdown disorder of the anthurium spathe.
Ca silicate gave slightly better results than Ca nitrate.
However, both were superior to "No treatment".

Experimental results show that the lack of Ca in the
anthurium is the major cause of the color-breakdown disorder
of the spathe and not oedema as originally suspected. The
lower Ca content in the lobe of the spathe as compared to
the tip section explains why the symptoms of the color-
breakdown disorder always occurs in the lobe of the spathe.
The pH and Ca uptake studies showed highest Ca uptake at
pH 7, lower at pH 5 and pH 9 and lowest at pH 3. This pH
factor coupled with environmental conditions of high
moisture and low temperatures could tend toward the
disorder in the winter months even though it did not appear
under experimental conditions. The benefit of applying Ca
to correct the disorder is twofold: 1) Ca is provided to
the plants as a nutrient and 2) the pH level of the medium
is raised to increase the chemical availability of Ca for the
plants.

A critical level of Ca within the tissue of the spathe
may be involved in the disorder. This critical level may be
very narrow but must be exceeded if tissue does not show
color-breakdown symptoms. This critical level may be
mediated by environmental conditions such as high air and

soil moisture, high soil and low air temperatures. The

95

Table 5. Field tests of the incidence of color-breakdown
disorder of spathe in anthurium with different Ca sources.

 

No. of flowers No. of flowers % flowers

 

 

 

 

Ca treatment w/o colorbreak w/colorbreak w/colorbreak
No treatment 2390 231 9.7
CA Nitrate

2000#/A/yr. 2640 32 1.2

Ca Silicate 2400 0 0.0

96

soil moisture, high soil and low air temperatures. The

hypothesis can be shown as:

 

 

Ca content in Environmental Expression of
regard to critical conditions that colorbreak symp—
level in the favor color- toms on the
tissue breakdown anthurium spathe

l. + - -

2. + + -

3. - - —

4. - + +

The color-breakdown disorder will mainfest itself on
the spathe only with the combination of weather and Ca as
indicated in #4, where the Ca level in the tissue is below
the critical level and the environmental conditions are
favorable for the color breakdown disorder. This explains
the higher incidence of the color-breakdown disorder as
reported by growers during the colder, rainy winter months
conditions favorable for color-breakdown as compared to
the warmer, sunny months conditions not favorable for color-
breakdown. The color-breakdown disorder has also been
reported to occur and then disappear on the same spathe
with variable weather conditions. This hypothesis would
adequately allow for such a phenomena.

Like the oedema disorder which is correlated with
environmental conditions which hinder the normal transpira-
tion processes of the plant, the anthurium color-breakdown
disorder may similarly be related to transpiration. Unlike

oedema, however, the critical level of Ca in the lobe of

97

the spathe is the major factor in this anthurium color-

breakdown disorder.

LITERATURE CITED

98

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Aragaki, M. 1973. Colletochrichum gloeosporioides
on anthurium spathe. Personal communication.

 

Balge, R. J., B. Esther Stuckmeyer and G. E. Beck.
1969. Occurrence, severity and nature of oedema in
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Chupp, C. and A. F. Sherf. 1960. Vegetable diseases
and their control. The Ronald Press Co. N. Y.
pg. 280-282.

Dale, Elizabeth. 1901. Investigation on the abnormal
outgrowth or intumescence on Hibiscus vitifoluis L.
Phil. Trans. Roy. Soc. of London. 194: 163-182.

 

 

Douglas, Gertude. 1907. The formation of intumes-
cenes on potato plants. Bot. Gaz. XLII(4): 233-250.

Eisa, H. M. and A. K. Dobrenz. 1971. Morphological
and anatomical aspects of oedema in eggplants. J.
Amer. Soc. Hort. Sci. 96(6):766-769.

Estabrooks, D. N. and H. Tiessen. 1972. Blossom-end
rot and black seeds of tomatoes. Can. J. Plant Sci.
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Ficq, A. 1959. Autoradiography, pp. 67-90. The cell,
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N. Y.

Geraldson, C. M. 1954. The control of blackheart
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Heggestad, H. E. 1945. Varietal variation and
inheritance studies on natural water soaking in
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Horwitz, W. 1960. Official methods of analysis of
the association of offical agricultural chemists.
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pg. 16 and 109.

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Jenkins, J., Jr. 1959. Brown rib of lettuce. Proc.
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Jensen, W. A. 1962. Botanical histochemistry. W. H.
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Kuster, E. 1903. Ueber experimentell erzeugte
intumeszensen. Ber. Deutsch. Bot. Gesells. 21: 452-
458.

Laganiere, J. and W. E. Sackston. 1967. Effects on
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Lange, A. H. 1959. Effects of environment on the
uptake-transport of calcium and phosphorus by bean
plants. Proc. Amer. Soc. Hort. Sci. 73: 349-354.

Lanning, F. C. and T. Garabedian. 1963. Distribution
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Lundegardh, H. 1935. The influence of the soil on the
growth of plants. Soil Sci. 40:423-437.

Maynard, D. N. and A. V. Barker. 1972. Internal
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102

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133: 64-69.

APPENDIX

CORRELATION OF THE ANATOMICAL STUDY
AND THE SPATHE COLOR-BREAKDOWN DISORDER OF

ANTHURIUM ANDREANUM, L.

 

103

APPENDIX

Correlation of the Anatomical Study and the Spathe

Color-Breakdown Disorder of Anthurium andreanum, L.

 

The following study was conducted to examine the
effects of the spathe color-breakdown disorder on the
internal anatomy of the anthurium flower.

Fresh anthurium tissues were taken of the spathe,
spadix and pedicel of plants showing and not showing
color-breakdown symptoms. The spathe tissues were
separated into lobe and tip sections. The tissues were
killed and fixed in FAA and histological procedures out-
lined by Jensen (1) and discussed under Materials and
Methods, Section I was followed. The tissues were studied
with an Olympus compound microsc0pe and pertinent data
recorded. Photomicrographs were taken with a Zeiss photo-
microsc0pe.

The cells of the color-breakdown spathe tissue were
collapsed. The collapsing was more intense in the mesophyll
than the epidermis (Figure 1). This is in accord with
earlier findings where X-ray analysis showed concentration
of Ca in the color-breakdown spathe tissue to be localized
in the epidermal areas and not in the mesophyll. The

color-breakdown tissues of the spathe were found to be

104

Figure 1.

105

A color-breakdown and a normal anthurium spathe
tissue. A=cross section of a normal spathe

tissue, photomicrograph, X16; E=cross section of

a color-breakdown spathe tissue showing the collapse
mesophyll tissue, photomicrograph, X16.

‘_——‘-—w—-—‘_--_P——‘-V——VV_'

/

. I . I:

a .iat..'u. n, Kw.

. \r . . x . I.) 'IV. . ,1 .9.

rant. Whiting 1&4 “Nut #‘m’ an...
- . . 1 - f.

55.... v
. £I...).I 61...: .L‘. 415.»...

C “a I J: r a“ g“ 4 1 . xl‘AMLAI
< . -. f -. t. a.

 

-..J.

107

torn and extremely difficult to find perfect, untorn
tissues. This was not true with normal tissues. The
tearing of the cells occurred between cells (e.g., separa-
tion of the middle lamella) and rarely through the cells
(Figure 2). This is similar to reports by Rasmussen (2)

in his studies with tender tobacco leaves. While the
tearing may have occurred during preparation procedures of
the tissues and not in the intact spathe, it indicates lack
of calcium pectate which results in the disintegration of
the middle lamella and separation of the cell wall. Studies
of the spadix and pedicel showed no sign of this primary
cell wall disintegration or separation. No difference in
anatomical structure between lobe and tip sections of
color-breakdown spathe could be found.

Differences in cellular details were difficult to
detect, as the protoplasmic constituents were not dis-
tinguishable with the histological procedures utilized.
Cell outlines and total protoplasmic materials, however,
were readily visible. Cells of the color-breakdown tissues
were found to take up less stain than cells of the normal
tissues. This was true in the spathe, spadix and pedicel.
This phenomena indicates the disintegration of the cellular
membrane (e.g., plasmalemma, tonoplast and the nuclear
membrane) and with the resulting internal leakage, a lack
of cytoplasmic materials.

No sign of cell hypertrophy or formation of intumes-

cense as found oedema tissue could be detected nor difference

 

Figure 2.

108

Color-breakdown anthurium spathe showing cellular
separation. Photomicrograph, X16.

 

 

 

 

110

in Ca crystal content in affected and normal tissues.

It is evident that more study on the cellular
details and the effect of low Ca must be pursued to com-
pletely comprehend the role of Ca in the color-breakdown
disorder. Tissue damage of the magnitude of cell separa—
tion and primary cell wall disintegration to cause this
disorder appears and disappears with changes in environmental
conditions. Ca is essential for the maintenance and for
the formation of cell-membrane systems on which the proper
functional integrity of cell metabolism is dependent, and
the lack of which has an effect on the internal chemistry
and on color change seems a more logical explanation. The
chemistry of the color change could be dependent on
environmental factors such as moisture in the cell and
manifest itself accordingly. Extreme cell damage leads
to irreversible breakdown and eventual necrosis of the
tissue. Ca effects on the primary cell wall are probably

secondary to the above mentioned changes.

I'l' l1l!n’il‘l

 

LITERATURE CITED

lll

LITERATURE CITED

Jensen, W. A. 1962. Botanical histochemistry.
W. H. Freemand and Company. S. E., California.

Rasmussen, H. P. 1967. Calcium andstrength of

leaves. I. Anatomy and histochemistry. Bot. Gaz.
128(3-4): 219-223.

112

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