1. INTEKNATIONAL RbVlEW OF CYTOLOGY. VOL. XV
Development of the Cotton Fiber
AMARJIT
S. BASRAAND C. P. MALIK
Department OJ Boticny, Pun,jub Agricultural Universily. Ludhiana, India
I.
11.
111.
IV.
V.
v1.
VII.
VIII.
IX.
X.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Growth Kinetics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Cytology of Early Fiber Development .........................
Chemical Changes during Fiber Development. . . .
Hormonal Considerations ...................................
Nutrients and Metabolites in Relation to Fiber Development. . . . . . .
Respiratory Changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Dark Metabolism of Carbon Dioxide.. . . . .
Cell Walls and Cell Wall Metabolism .........................
Some Concluding Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References
65
66
69
80
82
87
89
92
96
108
109
1. Introduction
Cotton is white gold. It is used for a variety of purposes, but especially to
make textiles used in the manufacture of a large proportion of man’s clothing.
With increasing world population the demand for cotton continues to increase
especially for light weight clothing fabrics in the tropics. Cotton is predominant
as a textile fiber because, as they dry, the mature fibers twist in such a way that
fine, strong threads can be spun from them. Cotton fibers are single-celled
outgrowths from individual epidermal cells on the outer integument of the ovules
in the developing cotton fruit. In the apt botanical expression, the fiber is a hair
or a trichome. The cotton fibers undergo a striking amount of elongation during
their development and can end up over 1000 to 3000 times longer than their
diameter. In the form of a single-celled epidermal hair, the cotton plant produces
one of the purest forms of cellulose known to man. The utilization of cotton
fibers dates back 7000 or 8000 years and fabrics woven from cotton are known
from 900 to 200 BC (Macneish, 1964). With such an antiquity, the fiber has
maintained its pristine purity and importance to this day.
In addition to its commercial importance, the developing cotton fiber has
several attributes that recommend it as an experimental system of choice for
investigation of physiological and biochemical changes accompanying cell
elongation and/or maturation. The fiber originates and ends as a single cell and
thus elongation can be studied free of any complications from cell division.
65

2. 66 AMARJIT S . BASRA AND C. P. MALIK
Fibers affirm a precise synchrony and homogeneity in growth during their devel-
opment in cotton bolls. They can be readily detached from the seeds and suffi-
cient material can be obtained for experimentation. The two rather distinct
phases of primary wall and secondary wall growth make the cotton fiber es-
pecially suitable for cell wall studies. Further potential advantages are that ovules
in a defined culture medium undergo normal morphogenesis including fiber
production (Beasley, 1977a; Kosmidou-DemCtrepoulou, 1979). The in v i m
methodology lends itself to detailed investigation of the factors that influence
fiber growth by exposing the ovules to various combinations of nutrients, metab-
olites, and phytohorniones. In this backdrop, the cotton fiber becomes a pivotal
plant structure to seek both fundamental and applied information.
In recent years, knowledge of the development of cotton fiber has shown an
impressive increase and some of the most significant research concerning cell
growth and cellulose biosynthesis has been carried out with this system. Except
for certain aspects (Flint, 1950;O’Kelley and Can, 1953;Beasley et nl., 1974b;
Beasley, 1977a;Berlinand Woodworth, 1980;Kosmidou-DemCtrepoulou, 1980;
De Langhe, 1980) no integrated review on this subject has appeared so far. The
purpose of this article is to review, summarize and evaluate various facets of
cotton fiber development and to focus attention on some currently critical areas
of investigation. Our hope is that this review will serve as a source of current
information to researchers in the field, but equally important will enable the
studentsof plant cell growth to become better informed about the interesting and
unique vistas which this plant structure provides for such studies. Comparison is
made with other experimental systems at places when it is considered appropriate
to explain the problem further.
11. Growth Kinetics
The seeds of the cultivated cottons bear relatively long hairs of commercial
importance, called lint or fibers and much shorter hairs called “linters” or fuzz
that have little commercial value. In the cotton trade, lint refers to those spinna-
ble fibers that are removed from the seed coat during the first pass through the
gin saws. The fuzz fibers remain adhered to the seeds. In the account that
follows, the discussion is confined mainly to the lint fibers.
On the basis of growth analysis, cotton fiber development has been divided
into four phases: ( I ) initiation, (2) elongation, (3)secondary thickening, and (4)
maturation (Naithani eta/., 1982).The fiber initiation starts a day before up to a
day or two after anthesis and the initials enter into elongation immediately. The
final length of a cotton fiber is the product of the rate of elongation per day and
the total period of elongation which is a genetic attribute (Fig. 1). The length ot

3. DEVELOPMENT OF THE COTTON FIBER 67
DAYS AFTER ANTHESIS
FIG.I . Rate curves of fiber length and dry weight against boll age in different cultivars of
cotton. (A) Gossypiurn hirsururn L. cv. Gujarat-67, (9)
G. hirsuturn L. cv. Khandwa-2, (C) G.
herbaceurn L. cv. Digvijay. (After Fig. 2 from Naithani e t a / . , 1982.)
the fiber largely determines the quality of the resulting thread. Variability in the
rate and the period of elongation and secondary wall deposition exists among
different cotton varieties. Older literature (Balls, 1915, 1928; Hawkins and Ser-
viss, 1930; Anderson and Kerr, 1938; O’Kelley and Carr, 1953; Kerr, 1966)
presented the concept that the secondary thickening phase does not begin unless
the elongation phase is completed. More recent and definitive work (Benedict et
al., 1973; Schubert et al., 1973, 1976; Meinert and Delmer, 1977; Beasley,
1979; Naithani et al., 1982) has shown a considerable overlap between the
elongation and the secondary thickening phases. The mechanism for coordinat-
ing the elongation and secondary thickening phases simultaneously in a develop-
ing fiber remains to be established. It may be possible to alter through genetic
manipulation either elongation or dry weight increase of the fibers without appre-
ciably changing the other (Kohel er al., 1974). Thus, fiber elongation and sec-
ondary wall deposition are not necessarily controlled by the same genetic factors.
There seem to be differences among different cotton varieties in the extent of
elongation which occurs after the onset of secondary wall formation (Beasley,
1979). Secondary wall cellulose deposition in fibers begins very sharply in
advance of cessation of elongation at a time related closely to final fiber length.
Although cause and effect relationships are not established yet it is possible that
onset of secondary wall elongation controls fiber length even though elongation
continues beyond the point of beginning of secondary wall thickening (Beasley,

4. 68 AMARJIT S. BASRA AND C. P. MALIK
1979). As the deposition of secondary wall does not immediately stop fiber
elongation, some specific wall thickness may be required to stop elongation
completely. Elongation occurs throughout the length of the fiber, not just at the
tip, although growth may be more rapid at the tip (Ryser, 1977; Willison and
Brown, 1977). According to Meinert and Delmer (1977), the fibers at early
stages of secondary wall deposition may elongate solely by the tip growth or the
cellulosic microfibrils may continue to be deposited in a random or transverse
orientation until the distinct increase in birefringence typical of secondary wall
appears. In view of the commercial desirability of long fibers, the study of
factors involved in controlling the extent of fiber growth is important.
Mature fibers exhibit thickened secondary walls composed of about 94%
cellulose and spiral twisting. The degree of thickening and the angle of spirals
affect fiber strength. Concomitant with the time of fiber maturity, the ovary wall
splits and opens along locular suture lines, leaving seeds and fibers exposed.
With the opening of the boll, loss of water and collapse of fiber cells occur and
the lumen contents dry into a residue.
The quality of fibers is important to the spinning and weaving industry and
determines the use to which it is put, as well as influencing the price paid for the
crop. Several characters are used to assess quality, some of them requiring
sophisticated measuring and testing devices. Staple length is the average length
of the fibers on a seed. It is an inherited characteristic of cultivars, which are
classified into five groups from “short” staple Asian cottons (less than 21 mrn)
to “extra long” staple Egyptian and Sea Island cottons (35 mm and longer).
Some 80% of world production is of “medium” (22-25 nim) and “medium
long” (25-28 mm) staple from Upland cultivars. The maturity of the fiber is
determined by the degree of secondary wall thickening laid down before it is
picked and, therefore, depends largely upon the time of crop harvesting. Fully
mature fibers have thick walls and a narrow lumen; they are strong and spin well.
On the other hand, immature fibers are not twisted and do not cling together
when the fiber is spun. Consequently, they produce tangles and knots of fibers
called “neps” in the yarn and “neppiness” in the cloth woven from it. Fiber
with high tensile strength is desirable because it is less liable to break during
ginning and spinning and because it produces strong yarn. Fine fibers with a
small diameter and fully developed walls are desirable because they produce the
strongest yarn for a given staple length. Good quality cotton, therefore, consists
of long, fine, and strong fibers.
Fiber length and fiber strength are greatly influenced by environment and
environment-genotype interactions (O’Kelley and Carr, 1953; Gipson and
Johani, 1968, 1969, 1970;Gipson and Ray, 1969;Quisenberry and Kohel, 1975;
Leffler, 1976; Kamsey, 1980). It is suggestive that fiber growth analysis in terms
of length and dry weight determinations on fiber from bolls of appropriate ages
may be useful in screening cotton lines for adaptability to certain environments.

5. DEVELOPMENT OF THE COTTON F I B I ~ R 69
111. Cytology of Early Fiber Development
The outer epidermal layer of the developing cotton ovule is composed of
epidermal cells, guard cells with subsidiary cells, and cotton fibers. The fibers
receive nutrition from the outer pigmented layer of the seed-coat underlying the
epidermis. This layer is several cells in thickness and is supplied with vascular
tissue. The foot of the fiber is absorptive in function (Fryxell, 1963). The early
development of fibers consists of two intergrading steps, which may be desig-
nated as spherical expansion above the ovular epidermis and elongation (Stewart,
1975). The epidermal cells are closely packed, cuboidal, and rich in cytoplasm
containing a large nucleus (Joshi eta/., 1967).The morphological differentiation
of a fiber begins when an epidermal cell rounds up and protrudes, the external
surfaces stretch outward, and the cell “baloons” above the epidermal surface
(Fig. 2). After the cells are fully expanded, the transition to elongation phase
begins. Once elongation has begun, the fiber cells do not divide. The transition
to elongation phase starts slowly as the blunt tipped cells begin to elongate
toward the micropylar end (Beasley, 1975; Stewart, 1975). During the second
and third day following anthesis, the rate of elongation increases and the fibers
segregate into groups. The rate of longitudinal growth apparently exceeds the
rate of diametric expansion as the tips become tapered. At this stage the fibers
also show spiral growth and no longer grow toward the micropyle. The stimulus
for directional growth of fibers toward the micropyle during the initial stages
remains obscure. The fiber surface is coated with a lamellar cuticle which
stretches and thins as the fiber elongates (Flint, 1950; Willison and Brown,
1977).
Although all epidermal cells (except the stomata1guard cells and cells com-
prising the micropyle) are potential fibers, not all differentiate into fiber initials
(Balls, 1915; Turner, 1929; Lang, 1938; Aiyangar, 1951; Joshi et al., 1967;
Beasley, 1975;Stewart, 1975). Fiber density is about 3300 per mm2and the ratio
of fiber-forming cells to the total number of epidermal cells is about 1 to 3.7 at
anthesis and the fibers do not occur in a regular pattern (Beasley, 1975;Stewart,
1975). This observation prompted research on cotton ovule culture with an
objective of increasing fiber yield per seed via pragmatically judicious and pre-
cisely timed applications of growth regulators (Beasley, 1977a). However,
events that determine which epidermal cells will differentiate into cotton fibers
remain to be established. Histochemical approaches have a vast scope to yield
significant qualitative information on the metabolism occurring in fiber-forming
and non-fiber-forming epidermal cells. The presence of stomata on cotton ovules
(Balls, 1919; Barritt, 1929; Aiyangar, 1948; Joshi et af.,1967; Elmore, 1973;
Beasley, 1975; Stewart, 1975) is also physiologically intriguing.
Both lint and fuzz fibers originate as epidermal outgrowths (fiber initials) of
the ovule. The fibers that initiate elongation on the day of anthesis are destined to

6. 70 AMARJIT S. BASRA AND C. P. MALlK
t
FIG 2 Dctdila of fiber initidtion (A) Ovule surtdce immediately before fiber initidtion Except
for stomdtd(S), differcntidting cells are not evident X82 (€3) Fiber initials ds they round up and
begin to expdnd No distinct pdttem of initiation IS evident X211 (C) Laterally cxpdnding fiber
initials Note fiber cell didmeter in reldtion to other epidermdl cells X166 (D) Fiber initials 1 day
dfter anthesis Elongdtion of thc fibers 15 toward the micropylar end (direction of anow) x50.5 (E)
Fiber initidh in all stages of development dt the rnicropylar end of an ovule 4 ddys dfter antheais
XI000 (Aftcr Figs 13-17, from Stewart. 1975 )
become lint whereas epidermal cells initiating elongation in subsequent waves
through about the fourth to twelfth day after anthesis, only develop into fuzz
fibers (Joshi et aZ., 1967; Beasley, 1977a). Both the range in time of initiation
and the extent of fuzz formed vary between species and cultivars. The phys-
iological and biochemicalbasis of lint and fuzz fiber formation is not understood.
Light and electron microscopy of fibers does not indicate a uniform prolifera-
tion of fibers over the whole seed. Light microscopy reveals that certain mor-
phological changes associated with the fiber differentiation occur at the chalaza1
end of the ovule 16 hours preanthesis and that additional cells undergo differ-

7. DEVELOPMENT OF THE COTTON FIBER 71
entiation closer to the micropylar end by 10 to 12 hours preanthesis (Aiyangar,
1951). At the ultrastructural level, initiation of fiber growth from cuboidal epi-
dermal cells is discernible between 24 and 16 hours preanthesis (Ramsey and
Berlin, 1976a). At 16hours preanthesis, differentiation of fiber initials is observ-
able at the chalaza. Occasionally, the fiber initiation from the crest of the
funiculus is noticed (Beasley, 1975; Stewart, 1975) (Fig. 3).
Fine structural alterations associated with early stages of fiber elongation
occur rapidly following anthesis and appear to be correlated with the formation
of the central vacuole, the plasma membrane, and the primary cell wall as well as
with increased protein synthesis necessary for extensive cell elongation (Ramsey
and Berlin, 1976b) (Figs. 4-7). A dilated portion of endoplasmic reticulum in
close association with the tonoplast showing a highly fenestrated membranous
network suggests the derivation of tonoplast of the central vacuole from the
endoplasmic reticulum. Formation of the large central vacuole begins at the base
of the fiber in a very precise manner and occupies most of the cell volume by 2
days after anthesis. Thus, a thin rim of cytoplasm separates the vacuole and the
cell wall during elongation, and the various organelles including the nucleus are
concentrated in the fiber tip. Dictyosome involvement in both plasma membrane
and primary cell wall formation is suggested from observations of similarities
between dictyosome-associated vesicles, containing fibrils appearing similar in
morphology to fibrils found in primary cell wall and plasma membrane associ-
ated vesicles. The differentiating fiber cells are enlarged and possess an enlarged
nucleus which is transposed from the original central position it occupies in the
ovular epidermal cell and an electron-dense cytoplasm due to the release of
phenolic substances from the vacuole and due to an increased number of ribo-
somes present in elongating fibers at anthesis (Figs. 8-10). Phenolic type com-
pounds have been observed to be 0-diphenols that presumably inhibit 1AA
oxidase to allow an intracellular auxin level high enough to initiate fiber differ-
entiation. In nondifferentiating fiber cells, the phenols are retained in the vac-
uole. Similarly, a growth-stimulating phenolic compound which stimulates the
process of fiber differentiation has been detected (Popova et al., 1979). More
numerous ribosomes and rough endoplasmic reticulum observed in fiber cells
than in adjacent nondifferentiating epidermis suggest a greater capacity for pro-
tein synthesis (Ramsey and Berlin, 1976a). The single nucleolus found in cotton
fibers enlarges following anthesis shows segregation of granular and fibrillar
components by 1 day after anthesis, develops a large “vacuole” thus appearing
ring shaped, and occupies much of the nuclear volume by 2 days after anthesis
(Ramsey and Berlin, 1976a). The fibrillar component is the first to receive newly
synthesized RNA which later passes to the granular component. Nucleolar vac-
uoles are spherical inclusions of low density, which characterize active nucleoli
and have roles in RNA transport. Nucleolar vacuolation during fiber growth
indicates simultaneous output and neosynthesis of nucleolar material (De Langhe

8. Fici. 3. Ovules at anthcsis, showing site and sequence of fibcr initiation on surface. (A) Crest of
funiculus (arrow) where fiber initials first appear. X84. (B) Side of ovule with arrow showing the
direction of progressive fiber initiation around the lateral circumference. XS I . (C) Chalaza1 end,
showing delayed fiber initiation at the tip. Note the numerous stomata. X72. (D)
Ovule with fibers
initiated in all area5 except the micropylar end (arrow). XSI. (E) Micropylar region of an ovule 4
days after anthesis with fiber initials (arrows). X500. (F) Lateral surface of ovule at anthesis. The
ratio of fiber initials to total epidermal cells is about 1:3.7, with about 3300 fibers per inm2 of
surface. Line represents 0. I X 202 mrn. (After Figs. 7- 12, from Stewart, 1975.)
72

9. FIG.4. Cotton fibers on the day of anthesis, the day elongation is initiated. (A) A median
longitudinal section of epidermal cells and an elongating fiber. The nucleus (N), possessing a single
nucleolus and a small amount of heterochromatin, has migrated toward the fiber tip. The first
manifestation of the central vacuole (V), containing both particulate and diffuse electron-dense
material, is apparent at the base of elongating cells. X7000. (B) Strands of endoplasmic reticulum
(E), continuous with the outer membrane of the nuclear envelope (arrow). are greatly dilated in
regions where the cisternae are closely associated with vacuoles (V) and with mcrnbranous networks
at the periphery of the vacuoles. Electron-dense particulate material is characteristically present in
these vacuoles. X44,OOO. (After Figs. 1-2, from Ramsey and Berlin, 1976b.)
73

10. 74 AMARJIT S. BASRA AND C P MA1.1K
FIG.5. Cotton fibers at 1 and 2 days after anthesis, respectively. (A) The enlarged central
vacuole (V) has moved out into the mid-region of the fiber with the base of the vacuole occupying a
position near the ovule surface. Some electron-dense material remains in the basal portion of the
vacuole, x4600. (B) The central vacuole (V) extended into the base of the fibers, leaving a thin rim
of cytoplasm in the fiber base and along with the mid-region of the fiber. X5000. (After Figs. 3-4,
from Ramsey and Berlin, 1976b.)

11. DEVELOPMENT OF THE COTTON FIBER 75
FIG.6. The fiber tip at 2 days after anthesis. (A) The tip of elongating fibers is filled with
cytoplasm containing numerous dictyosomes (D), ribosomes, strands of endoplasmic reticulum (E),
mitochondria (M), small vacuoles (V), and lipid bodies (L). X 14,500. (B) An increased magnifica-
tion of a portion of A shows dictyosomes ( 0 ) with electron-dense mature-face cisternae and fibrils
(arrows) in the dictyosome vesicles. X56,500. (After Figs. 5-6, from Ramsey and Berlin, 1976b.)

12. FIG.7 . The cytoplasm of the mid-region of young cotton fibers. ( A ) Portions of 2 cotton fihers at
2 days after anthesis reveal similarities between dictyosomal and plasma membrane associated
vesicles (arrows). Both types of vesicles are similar in size and both contain fibrils morphologically
similar to fibrils found in the primary cell wall (W). X28,OOO. (B) A ph5nla menlbrdne elaboration
showing an electron-dense membrane and fibrillar contents. x46.000. (C) A plasma membrane
elaboration. showing continuity between the interior of the elaboration and the primary cell wall (W),
is apparent. ~76,000.
(After Figs. 7-9, from Ramsey and Berlin, 1976b.)

13. DEVELOPMENT OF THE COTTON F:IBIiR 77
FIG.8. Ovule epidermal cells. (A) Epidermal cells at the chalaza1 end of ovule at 16 hours
preanthesis; most of electron-dense material has bcen dispersed from vacuoles (V) of these three cells
giving them a dark appearance (compare with subepidermal cells) X 10,000, (B) An early stage in
release of phenolic compounds in an epidermal cell at antheais is indicative of fiber differentiation.
Cytoplasm between the vacuole (V) and the nucleus (Nu) is more electron dense than cytoplasm in
the other parts of cells apparently due to adherence of phenolic material to the cytoplasmic or-
ganelles. X 12.000. (After Figs. 5-6. from Ramsey and Berlin, 1976a.)

14. FIG.9. Comparison of light and dark epidermal cells. (A) Median longitudinal section of ovule
epidermal cells approximately one-third of ovule length from chalaza1 end showing electron-dense
“cldrk” and less electron-dense “light” epidermal cells at 8 hours preanthesis. X 10,000, (B)
Portions of a nonelongating light cell (L) and a differentiating dark primordial fiber cell (D) at
anthesis. Dark cell has an electron-dense layer of material on the cytoplasmic surface of the plasma
membrane (PM), endoplasmic reticulum (E). ribosomes (unlabeled arrows), and mitochondria (M).
This dense layer is absent at the membrane surface away from the cytoplasm, for example, the
plasma membrane of dark cell adjacent to the cell wall (W). X65.000. (C) Portions of a light (L) and
dark (D) cell at anthcsis. Electron-dense material in dark cell coats the cytoplasniic surfaces of the
plasma membrane (PM). endoplasmic reticulum (ER), ribosomes (unlabeled arrows). and mito-
chondria (MI.
The asymmetric deposition of electron-dense material is apparent on cell wall (W)face
of plasma membranc of dark cell. X80,OOO. (After Figs. 8-10 from Ramsey and Berlin, 1976a.)
78

15. FIG. 10. Differentiating fiber cclls iniriating elongation on the day of anthesis. (A) A dark
epidermal cell is protruding above ovular surface. Endoplasmic reticulum (E) is well developed in
primordial fiber cell. Only a small amount of electron-dense material remains in most vacuoles (V).
Note light cells on either side of dark cell. X 12,000. (B) An elongated fiber cell with enlarged
nucleus (Nu) and nucleolus. Nuclcus has migrated toward fiber tip. Note division in epidermal cell
adjacent to fiber cell. ~ 5 0 0 0 .
(C) Scanning electron micrograph of ovule surface showing lint fibers
protruding above the epidermal surface. XSOO. (After Figs. 11-13, from Ramsey and Berlin,
1976a.)
79

16. 80 AMARJIT S. BASKA AND C.P. MALlK
et a/., 1978). Prominent nucleoli are not observed in nuclei after 10 days of
anthesis, suggesting that ribosome synthesis necessary for fiber development
occurs early in the elongation period which declines significantly during later
stages of elongation and maturation (Ramsey and Berlin, 1976b). Thus, the
amount of ribosomes synthesized in very early stages of fiber elongation may
subsequently determine the rate of elongation and thickness of the fiber as well
(Rarnsey and Berlin, 1976a,b; De Langhe rt NI., 1978).
IV. Chemical Changes during Fiber Development
Mature cotton fiber contains on an average (percentage of absolute dry sub-
stance) cellulose, 94.0; wax-like substances, 0.6; pectins (calculated as pectic
acid), 0.9; organic acids, 0.8; nitrogenous substances (calculated as proteins),
1.3; ash, I .2; noncellulosic polysaccharides, 0.3;and, unidentified substances,
0.9 (total, 100’%).
The sooner the fiber is gathered, i.e., the smaller the degree of its maturity, the
lower the cellulose content and the greater the amount of other admixtures and
the moisture content. (Different aspects of cellulose will be dealt with later in
Section IX.) There is considerable synthesis of proteins during fiber development
(O’Kelley and Cam, 1953; Huwyler e
f uf., 1979). Ontogenetic changes in es-
terase and alkaline phosphatase have been investigated in developing fibers
(Rama Kao et d.,
1980; Rama Rao and Singh, 1982b). Early work showed that
the level of reducing sugars in fibers is appreciably high during the elongation
phase which decreases during secondary wall formation (O’Kelley and Carr,
1953). The glucose and sucrose are the only major ncutral sugars found in the
fibers (Carpita and Delmer, 1981). Cotton bolls accumulate good amounts of
carbohydrates, mainly glucose, fructose, and sucrose during developnient (Con-
ner et ul., 1972). Recently, Jaquet ef a/. (1982) have measured changes in these
sugars in individual fibers of Gossypilrm spp. at different stages of development.
The results indicate that during primary wall formation, sucrose which is the
transport sugar, is inverted and that glucose and fructose are accumulated for
later use in the synthesis of secondary cell wall. On the other hand, the sucrose
content increased regularly until fiber maturity. There exist separate “storage”
and “metabolic” pools of glucose in the fiber showing mutual exchange (Carpita
and Delmer, 198I ). Since the fibers are highly vacuolated, the storage pool is
probably the vacuole. This is supported by thc fact that only about I IYo of the
total reducing sugars of the cell is susceptible to rapid release by treatment of the
fibers with 7.5% dimethyl sulfoxide (Carpita and Delmer, 1981). Such treatment
has been shown to alter permeability of the plasma membrane while having much
less effect on the vacuolar membrane of the plant cells (Delmer, 1979).
Cotton bolls accumulate minerals throughout their development. Redistribu-

17. DEVELOPMENT OF THE COTTON FIBER 81
tion of minerals (nitrogen, phosphorus, and potassium) among bur, seed, and
fiber may account for many of the compositional changes in each component
especially during the period of boll maturation (Leffler and Tubertini, 1976).
This suggests the existence of a physiological continuum among the boll compo-
nents during development. Starch does not represent a major portion of the
carbon in fibers (Flint, 1950; Meinert and Delmer, 1977; Maltby et al., 1979).
Ascorbic acid content is high in young fibers and decreases at maturity (Jasdan-
wala et al., 1980). Analysis of organic acids from the fiber shows malate and
citrate to be the predominant ones (McCall and Guthrie, 1945; Dhindsa et al.,
1975). There is a measurable amount of water-soluble arabinogalactan-likepoly-
mers in the fibers (Carpita and Delmer, 1981).
Mature fibers contain 0.5%of a lipid which is a mixture of waxes, fats, and
resins (Amin and Truter, 1972;Ferretti et al., 1975; Iyengar et nl., 1982). The
lipid content in young fibers, however, is quite high. The fibers undergoing
active extension incorporate most of the label from [I-14C]acetate into polar
lipids as compared with the nonpolar lipids implying active membrane bio-
synthesis (Basra and Malik, I983a). Lipid synthesis contributes to the tonoplast
enlargement and thereby generation of turgor pressure. Apparently, the amount
of lipid synthesized during fiber growth functions mainly in the synthesis of
membranes and maintenance of their biochemical integrity. The presence of lipid
bodies, sterols, steryl glucosides, esterified steryl glucosides, glucosyl-phos-
phorylprenol, fatty hormones, etc. has been ascertained in developing fibers
(Mandava and Mitchell, 1971; Forsee and Elbein, 1972, 1975; Forsee et al.,
1974, 1976; Beasley, 1975; Ramsey and Berlin, 1976b; Delmer ef a/., 1977;
Carpita and Delmer, 1981). A particulate enzyme system from cotton fibers
forms both steryl glucosides and acylated steryl glucosides by catalyzing the
transfer of [ 14C]glucosefrom UDP-[14C]glucoseto endogenous sterol acceptors
(Forsee ef id., 1974). Analysis of the products by gas-liquid chromatography
and mass spectrophotometry revealed that p-sitosterol is the predominant sterol
moiety, while campesterol, cholesterol, and stigmasterol are present in smaller
amounts. Palmitate and oleate are the major acyl components of the esterified
glucoside. The appearance of radioactivity first in the steryl glucoside and then in
the acylated steryl glucoside suggests a precursor-product relationship whereby
the steryl glucoside is the immediate precursor of the acylated steryl glucoside
(Forsee rt al., 1974). That the steryl glucoside is indeed the precursor for the
acylated steryl glucoside has been shown to be the case by incubation of steryl
[14C]glucosidein the presence of particulate enzyme from fibers. As a function
of time, radioactivity disappears from glucoside and appears in the acylated
steryl glucoside (Forsee et al., 1976). The acyl transferase that is involved in the
transfer of acyl group to the steryl glucoside has been partially purified. Phos-
phatidylethanolaminehas been shown to be the best acyl donor by demonstrating
that 14C-labeledfatty acids from 14C-labeledphospholipid can be transferred to

18. 82 AMARJIT S. BASRA AND C. P. MALlK
steryl 13H]glucosideto form a I4C, 3H-labeled acylated steryl glucoside. The
steryl glucosides and their acylated derivatives are found in many membranesbut
the function of these compounds is unknown (Elbein, 1980). It has been sug-
gested that they may play a role in membrane permeability and that they may
have hormonal action.
Colorimetric determinations of proline and hydroxyproline in developing
fibers demonstrate their presence in the wall, proteins, and soluble fractions
(Basra, 1982). The study noticed that hydroxyproline content in fiber walls of a
short staple cultivar is higher than its long counterpart during the period of rapid
expansion (Basra, 1982). Although, the amount of hydroxyproline is low in the
cotton fiber cell walls, the presence of some 2 linked arabinosyl residues in these
preparations could indicate the existence of hydroxyproline arabinosides
(Meinert and Delmer, 1977). However, the absolute amounts of hydroxyproline
detected in the walls may not be the limiting factor for cell extension (Basile,
1979). It is probable that the degree to which the fibers will attain their final
length is contingent upon the time and mode of deposition and/or functional
relationshipof certain hydroxyproline containing protein(s) to other wall compo-
nents in elongating cotton fibers.
V. Hormonal Considerations
Considerable evidence indicates that hormones play a decisive role in fiber
development (Kosmidou-Dkmktrepoulou, 1980). Studies in this direction have
been facilitated to a great extent by the culture of both fertilized and unfertilized
cotton ovules (Fig. 11). For a detailed account, the reviews by Beasley (1973,
1977a) and Beasley et al. (1974b) are indispensable. The in vitro methodology
lends itself to a greater range of environmental and chemical manipulations than
are possible with the whole plants. The basal culture medium for cotton ovules is
listed in Table I. Total fiber development is assessed by the stain-destain method
(Beasleyet al., 1974a). Briefly, the method is as follows: (1) 20 ovules (all from
a single ovary) with associated fibers are placed for 15 seconds in 80 m
l of
0.018% toludine blue 0, (2) nonabsorbed dye is removed by a 60-second run-
ning-water wash, (3) absorbed dye is removed by 100 ml of destaining solutiQn
(1 part glacial acetic acid, 9 parts 95% ethanol), and (4)absorbance of destaining
solution is then determined after 1 hour of destaining. Absorbance values are
used as a measure of fiber development and are expressed in terms of total fiber
units (TFU); one 00 unit at 624 nm has been assigned the value of one TFU. Dry
weights of ovules and their associated fibers, pooled by treatments, are often
determined after recording TFU.
Fibers on isolated ovules continue to develop in culture, only if fertilization is
accomplished before harvest of the ovaries and transfer of ovules to a liquid

19. DEVELOPMENT OF THE COTTON FIBER 83
FIG. 11. Cultured ovules of cotton. (A) Fertilized ovules in liquid medium. Fibers have con-
tinued to elongate and embryos to develop normally, even to the point of germination. (B) Unfer-
tilized ovules from flowers in which fertilization has been prevented. Some ovules have enlarged
slightly in culture but no fibers have developed. (C) Unfertilized ovules cultured in medium contain-
ing indoleacetic acid and gibberellic acid. With the addition of these hormones unfertilized ovules
have enlarged and produced fibers (compare the fertilized ovules in A). (D) Unfertilized ovules from
(C) treated so as to extend the fibers. (After Fig. 2, from Beasley and Ting, 1974.)
growth medium (Beasley, 1971). GA, markedly promotes the total amount of
fiber produced from fertilized ovules (Beasley et al., 1971). The fertilized iso-
lated cotton ovules appear to be (1) deficient in their capacity to synthesize
optimal levels of gibberellins, (2) sufficient in their production of cytokinins, (3)
optimal or near optimal in the production of auxin (IAA), and (4) ABA is not
essential for fiber elongation and a diminution of its effective concentration

20. 84 AMARJIT S.BASRA AND C. P. MALIK
TABLE I
BASAL CLILTUKt Mt:I)ILIM FOR CUII'ON OVLILES"."
Stock giliter ml stockiliter ingiliter nuM
number Component (stock) (final) (final) (final)
27.2 I80
0.6183
0.0242
44.1060
0.0x30
0.0024
49.3000
I .6"2
0.8627
0.0025
505.5500'
0.8341
1.1167
0.0492
0.0822
0.I349
18.0I60
-
10 212 180
6 183
0 242
10 441 060
0 024
10 493 000
16 902
8 627
0 025
20 5055 500
10 8 341
II 167
I0 0 492
0 x22
1 349
10 1x0 I60
- 2 I620 000
n 830
2 0000
0 1000
0 0010
7 0000
0 0050
0 0001
2 0000
0 1000
0 0300
0 0001
50.0000
0 0300
0 0300
0 0040
0 0040
0 0040
I 0000
120 0000
"pH adjusted to 5.0 prior to autoclaving.
'>Formaximum fiber production from fertili7ed ovulcs, 0.5 -5.0pM tiA1 is used. For occasional
slight stimulation, 5.0 pM IAA ia also employed. For inaxiiiiuiii fiber production from unfertili7cJ
ovulcs, 5.0 pM IAA and 0.5 KM GA7 are employed. For occasional slighl stimulation, 0.05 pM
kinetin i$ also employed and/or KN03 is reduced to 45 mM and 2-5 mM NHJNOq is added. TWU
methods for the induction of callus fi-or11cultured ovulcs (unfertilized) arc subhtitutc tructose lor
glucose and cmploy 5.0 pM GA7,or use plastic cultured vessels, Jelctc boron, employ NH4N0,3as
in 3 abovc, and suhstitutc sucrose foi- glucose (after. Rcaslcy, 1977a)
~'Amounti2
I s t d (g).
"Ainbcr bottle.
<'Refrigerate.
concomitant with and perhaps dependent upon an increase in IAA and GA,
following fertilization permits ovular and fiber growth (Beasley and Ting, 1973).
The unfertilized ovules, on the other hand, require addition of IAA to the basal
medium in order to produce fiber (Beasley, 1973)and it is suggested that auxin is
the major hormone produced in response to the process of fertilization, whereas
gibberellins probably derived, for the niost part, from sources external to the
ovule (Beasley and Ting, 1974). Simultaneous additions of IAA and GA, pro-
duce additive amounts of fiber froin unfertilized ovules (Beasley and Ting,
1974). ABA reduces the amount of fiber produced by IAA and k;Aetin partially
overcomes the inhibition caused by ABA. The ovules acquire their capacity to
respond to phytohormones between the third and second day preanthesis.

21. DEVELOPMENT OF THE COTTON FIBER 85
Further, Birnbaum et al. (1974) report that IAA, whether synthesized endoge-
nously in fertilized ovules or added to the growth mcdium of unfertilized ovules,
may be of greater importance to fiber production than gibberellins. Dhindsa
(1978a) concluded that ( I ) in the presence of the antiauxin, PClB (p-chlo-
rophenoxyisobutyric acid) alone and in combination with GA,, or GA, + IAA,
unfertilized cotton ovules grow in size but do not produce fibers; and (2) ovule
growth appears to be predominantly determined by gibberellin while fiber
growth is largely dependent on the availability of auxin. It is also speculated that
(1) the generally agreed upon differences between time of initiation for fuzz and
lint fibers, and (2) the relatively distinct differences in length of the two fiber
types, are due to sequential “perception” and relative amounts of effective
endogenous auxins and gibberellins (Beasley, 1977a). Isolated fiber protoplasts
form new walls in culture and divide to form callus. Removal from the cell wall
thus leads to division rather than expansion of the protoplast (Beasley ef ul.,
1974b).
Isolation of plant growth substances from developing cotton fruit has been
reported (Davis et al., 1968; Sandstedt, 197I , 1974; De Langhe, 1973; Morgan
el ul., 1972; Shindy and Smith, 1975; Rodgers, 1981a-c; Guinn, 1982). Howev-
er, few investigators have studied these substances in individual fibers (Mitchell
et a/., 1967; Mandava and Mitchell, 1971; Naithani, et a/., 1982). The cotton
fibers are distinctive in having a special type of growth hormones which are
lipids containing fatty acids with 14 to 22 Carbon atonis (Mandava and Mitchell,
1971). The cotton fibers were found to be a rich source of auxin substances by
the bioassay method (Naithani e t a / ., 1982). In this study, the long staple cultivar
had the highest content of auxin substances followed by the medium and short
staple cultivars (Fig. 12). However, changes in auxin substances did never show
any correlation with the rate of fiber elongation and the peak levels of auxin
substances in all the cultivars were recorded before or about the time when
elongation had just started. In this connection, there is a general failure to
correlate levels of plant growth substances with the developmental event sup-
posedly being regulated (Trewavas, 1982; Hanson and Trewavas, 1982). The
sensitivity and reliability of the bioassay method are controversial, Therefore,
investigations with latcst tcchniques like combined gas chromatography-mass
spectrometry and radioimniunoassay should be carried out.
Using bioassay, Bhardwaj and Lad (1 977) investigated the endogenous levels
of auxins, gibberellins, and growth inhibitory substances in lintless and linted
genotypes of G. arhoreum L. The genotypic variation in fiber length was found
to be related with gibberellin content of the seeds. However, it was not clear
whether lintlessness was caused by the deficiency of gibberellins alone or im-
posed by accumulation of inhibitors in large amounts in the pericarp or both. A
one-time addition of GA, in situ to flowers emasculated before anthesis can
replace pollination by 100% for fruit wall growth and by some 50% for fiber
elongation (Baert et ul., 1975). Addition of auxins to these GA,-treated flowers

22. 86 AMARJIT S . BASRA AND C. P. MALIK
DAYS AFTER ANTHESIS
FIG. 12. Auxin content against boll age in different cultivars of cotton. (A) Goss!pirr,n hirsrctron
L. cv. Gujara(-67, (B) G. hirsrrfrtrrr L. cv. Khandwa-2. (C) G. hurhocrurn L. cv. Digvijay. (After
Fig. 3 . from Naithani pr ( I / , , 1982.)
stimulates further fiber elongation and the mature fibers obtained are comparable
to nornial fibers although secondary wall formation is generally less pronounced.
Indeterminate growth habit of the cotton plant almost precludes that the growth
substances [even if the right onc(s) are applied] are available at the required site
at the time the fiber might have been favorably responsive (Bcasley et a/..
1974b). However, thcrc exists a theoretical possibility that with the advent of
determinate cottons. the application of growth substances may prove to be effec-
tive in increasing the number, Icngth, thickness, and uniformity of cotton fibers.
Low activities of oxidative enzymes, IAA oxidase, peroxidase, and O-di-
phenol oxidase in preanthesis ovules and an increasing trend until 5 days after
anthesis, were rccordcd (Jasdanwala Pt a/., 1980;Kama Kao and Singh, 1982a).
On the day of anthesis, very low IAA oxidase was recorded indicating that IAA
is necessary for fiber initiation. It was suggcsted that a shift in redox balance
from a reduced state to an oxidative state in developing ovules results in fiber
initiation. The quite extensive work on IAA oxidase and peroxidase has shown
that auxin catabolism is low during the elongation phase but very high in the
secondary thickening period (Jasdanwala ct ol., 1977, 1980; Basra and Malik,
I98 I ; Rama Rao et a/., I982a,b). It is possible that the total period of elongation
is regulated by an auxin degrading system, as an increase in auxin degradation
might decrease the availability of auxin for elongation growth (Naithani rt cil.,
1982; Kama Kao et ctl., 1982a,b). Concomitantly, 0-diphenol oxidase activity is

23. DEVELOPMENT OF THE COTTON F i w K 87
low during the elongation phase but increases sharply during the secondary
thickening phase (Naithani et a/., I981). High 0-dihydroxyphenols and low
activities of IAA oxidase and peroxidase in elongating fibers are in accordance
with the cellular environments favorable for a rapid rate of cell growth.
It is noteworthy that the activities of IAA oxidase and peroxidase are consider-
ably higher during the elongation phase in the short staple cultivars than the
medium or long staple cultivars (Basra and Malik, 1981; Rama Rao er al.,
1982a,b). Thus, both the availability of IAA for growth and the total period of
elongation are reduced in the short staple cultivar (Rama Rao ef al., 1982a,b). In
line with this is the report that the auxin content is higher in the fibers of the long
staple cultivar than the short counterparts (Naithani et ul., 1982). Some of the
physiological control points for IAA oxidase and peroxidase catalyzed oxidation
of IAA are action and interaction of phytohormones, minerals, phenols, cou-
marins, organic acids, and redox regulators (Sembdner era/., 1980)which may
interact in sonic way regulating the enzyme levels of cotton fibers.
Peroxidase is a multifunctional enzyme which can regulate cell extension in
the capacity of an exocellular glycoenzyme (Lamport, 1980). The ionically
bound wall peroxidase activity kept low levels during the elongation phase and
high levels during the secondary thickening phase (Rama Kao et al., 1982b) and
the possibility of wall peroxidase in cessation of fiber growth was considered.
V1. Nutrients and Metabolites in Relation to Fiber Development
The in vitro methodology of cotton ovule culture has been used to study the
role of micronutrients in ovule and fiber growth. Maximum effort along these
lines has been expended on the role of boron in ovule and fiber growth. A
constant supply of boron is necessary to maintain fiber elongation and prevent
callusing of epidermal cells in vitro. In the boron-deficient medium, ovules
callus and accun~ulate
brown substances (Birnbaum ei al., 1974). Development
of boron deficiency symptoms in the cultured ovules is determined partly by the
phytohormones include& in the basal medium. Profuse callusing in the absence
of boron occurs only in the presence of GA,. Thiamine is the critical vitamin
essential for GA,-induced callus formation when unfertilized ovules are cultured
in the absence of boron (Birnbaum et ul., 1974). The studies led to the conclu-
sion that exogenous thiamine was not essential to the continued elongation of
fibers that had already initiated growth on the day of anthesis, but was essential
for growth of integuments (including epidermal cell divisions) and development
of new fiber initials beginning their elongation phase after the day of anthesis
(after transfer to culture). It seems unlikely that a thiamine deficiency alone
would be the cause of altcred ratios in lint to fuzz fibers seen among cotton
varieties (Beaslcy, 1977a). It was pointed out, however, that more sophisticated

24. 88 AMARJIT S . BASRA AND C. P. MALlK
experiments with thiamine might lead to information valuable in explaining the
physiological and biochemical basis for altered lint fuzz percentages.
Boron deficiency-like symptoms are induced by 6-azauracil (inhibitor of
orotidine monophosphate decarboxylase) in ovules growing in boron-sufficient
medium (Birnbaum et al., 1977). The correlation is further strengthened by the
finding that orotic acid and uracil partially overcome both boron deficiency and
azauracil effects. These studies suggest that boron deficiency symptoms are
related to reduced activity in the pyrimidine biosynthetic pathway. In this way,
boron deficiency may cause reduced synthesis of UDP-glucose and other UDP-
sugars involved in cell wall composition of the fiber. Wainright et (11. ( I 980)
sought to establish more directly whether boron regulates the pyrimidine path-
way in some way by studying incorporation of ['4CJoroticacid into intermedi-
ates of the pyrimidine pathway. Total incorporation of [6-14C]oroticacid into
fiber was inhibited by 59% under boron deficiency. The inhibition was evident in
all radioactively labeled pools, indicating that the effect may be at the membrane
transport level or at an early stage of orotic acid metabolism (e.g., inhibition of
orotodine monophosphate decarboxylase). In other plant systems, evidence is
accumulating that boron may play an important role in membrane transport or in
maintaining membrane integrity (Pollard et ul., 1977;Hirsch and Torrey, 1980;
Roth-Bejerano and Itai, 1981; Hirsch et ul., 1982).
The second major effect of boron deficiency in the in vitro cultured cotton
fiber system is the high percentage incorporation into RNA than under sufficien-
cy (Wainrightet al., 1980). Conversely, the percentage incorporation into UDP-
glucose is lower under boron deficiency. The incorporation of labeled UDP-
glucose into cell wall material of fibers is also reduced under boron deficiency
(Dugger and Palmer, 1980). There is evidence that UDP-glucose pyrophos-
phorylase has strong product inhibition (Gustafson and Gander, 1972; Hopper
and Dickinson, 1972). Therefore, if UDP-glucose were to accumulate due to
nonutilization in the synthesis of cell wall material, one would expect rapid
inhibition of UDP-glucosepyrophosphorylase. This would result in the observed
decreased radioactive orotic acid incorporation.
In sum, the studies indicate that boron deficiency .in cotton fibers causes a
general inhibition of orotic acid incorporation whereby pyrimidine synthesis
intermediatesare shunted away from UDP-glucosesynthesis and channeled pref-
erentially into RNA synthesis. This could be related directly to cessation of fiber
growth due to inhibition of wall synthesis. The experimental data just do not
exist at this time to make an overall cirtical assessment of the role of various
mineral ions in the control of fiber morphogenesis.
NH4+ is another important factor in the growth and development of cultured
cotton ovules (Beasley and Ting, 1974; Beasley, 1977b). For example, the
ovules cultured at 28°C require IAA and either NH4+ or GA, in the substratum
for fiber development whereas ovules cultured at 34°C require only IAA. NH, + ,

25. DEVELOPMENT OF THE COTTON F113ER 89
GA,, IAA, or increased temperature have no effect on the induction of increased
nitrate reductase activity in the ovules so that the effects of these compounds on
fiber development are independent of the availability of reduced nitrogen as a
general substrate for growth (Beasley et ul., 1979). Also, the ovules receive
reduced nitrogen almost exclusively in vivo (Radin and Sell, 1975). The mecha-
nisms of NH, + and high temperature in regulation of fiber development remain
elusive.
VII. Respiratory Changes
Cell growth depends upon metabolic energy and biosynthesis. During carbo-
hydrate oxidation, the energy stored in carbohydrate molecules is tapped for the
endergonic activities of cells. At the same time, the metabolism of carbohydrates
provides a number of intermediates for biosynthetic processes and cellular main-
tenance. It is probable that nonphotosynthetic and nongluconeogenic plant cells
receive the bulk of their organic carbon as sucrose (apRees, 1977)and hence the
breakdown of sucrose rather than the metabolism of hexose is the starting point
of carbohydrate metabolism in these cells.
Sucrose utilization is initiated by two enzymatic reactions: (1) hydrolytic
cleavage to D-glucose and D-fructose by the action of invertase, and/or (2)
cleavage by sucrose synthetase to produce fructose and sugar nucleotide inter-
mediates, primarily UDP-glucose. High activities of acid invertase and sucrose
synthetase in both in vivo and in vitro grown fibers have been reported (Beasley
et al., 1974b). High acid invertase ensures a large net demand of hexoses for
extensive fiber extension. Buchala and Meier (unpublished data) have demon-
strated the presence of cell surface located invertase for G. urboreum fibers and
one of its possible roles in regulation of cell wall synthesis by mediation of
uptake of sucrose from the apoplast into the symplast has been suggested. The
reaction of hexokinase funnels hexoses into intermediary metabolism by phos-
phorylating glucose and fructose to the corresponding hexose 6-phosphate (Turn-
er and Turner, 1980). The hexose monophosphates once produced may undergo
glycolysis, enter the pentose phosphate pathway, participate in oligo- or polysac-
charide synthesis, or be hydrolyzed to free hexoses by phosphatases. The hex-
okinases, particularly, those which display high activities toward D-fructose, are
most important in the metabolism of rapidly developing nonphotosynthetic cells
which have large requirements for precursors of cell wall polysaccharides
(Feingold and Avigad, 1980). The results of most studies are consistent with
provision of a major part of UDP-glucose in nonphotosynthetic cells by the
action of sucrose synthetase (Feingold and Avigad, 1980). UDP-glucose is en
route to the synthesis of cellulose in developing cotton fibers (Carpita and Del-
mer, 1981). Thus, a direct conversion of sucrose to sugar nucleotides for cell

26. 90 AMARJIT S.BASRA AND C. P. MALlK
wall synthesis could account for a significant fraction of sucrose breakdown in
cotton fibers. It is likely that the rate of sucrose import and growth of fibers may
be controllcd by an interplay of invertase and sucrose synthetase which ensure
that the rate of import is precisely matched with the rate of its metabolic utili-
zation.
It has been shown that glycolysis and the pentose phosphate pathway operate
in elongating cotton fibers and that the extent of their operation varies with the
demand for respiratory products (Basra, 1982). In this respect, hexokinase,
glucose-6-phosphate dehydrogenase, phosphofructokinase and pyruvate kinase,
and succinatc dehydrogenase show increased activities during the period of rapid
extension growth and decreased activities when the rate of growth slows down.
The oxidation of [ I-'JCJ- and [6-'4C]glucose and measurements of important
glycolytic and pentose phosphate pathway intermediates yielded a similar pat-
tern. The increased channeling of metabolized glucose into thc two pathways
during the period of active fiber growth reflects a high requirement for energy
and reducing power which must be produced to attain cell extension of consider-
able magnitude. Cotton fibers have high levels of adenosine phosphates and low
energy charge during this period (Basra and Malik, 1982). As the rate of fiber
growth slows down, the decline in enzyme activities, metabolites, and turnover
rates of [ 14C]glucosepoints to a shift in metabolic priorities. A large proportion
of the carbon budget at later stages of fiber development is expended in the
synthesis of a thick secondary wall. Mutasers (1976) indicated that over 60%of
the translocated sugar accumulates in wall polymers and Carpita and Dclmer
(198 I ) observed that a high proportion of the carbon passing through the metabo-
litc pool of glucose is used for the synthesis of ccllulose and p,I-3-glucan alone.
However, a significant level of glucose oxidation continues to support the on-
going fibcr development.
C&, ratios are less than unity during the period of fiber elongation and thus
provide strong evidence for the operation of pentose phosphate pathway (Basra,
1982). It is logical that the pentose phosphate pathway is functionally more
important in nonphotosynthetic cells to compensate for the NADPH, normally
produced in the chloroplasts. Biosynthesis usually requires NADPH as opposed to
NADH. The cotton fibers are equipped with an additional pathway of NADPH
generation. It is seen that when the activity of the pentose phosphate pathway
decreases during the later stagesof fiber growth, malate accumulatedvia CO, dark
fixation is catabolized to supply NADPH via decarboxylation by malic enzyme
(Basra and Malik, 1983h) and resulting thereby in the increase of C&, ratios.
Glucose-6-phosphate dehydrogenase is controlled in vivo by the NADP + /
NADPH ratio (apRees, 1980b). It is just conceivable that an increase in the
activity of the pentose phosphate pathway may reduce the rate of malate decarbox-
ylation or vice versa through the regulation of the NADP+/NADPH ratio.

27. DEVELOPMENT OF THE COTTON l ~ l l 3 l ~ K 91
ATP, ADP, and AMP have been identified in cotton fibers (Franz, 1969;
Carpita and Delmer, 1981) and determinations at different periods of fiber
elongation have been made (Basra and Malik, 1982). It has been proposed that
rate of fiber elongation may be the result of ATP levels (Basra and Malik, 1982).
One of the most important cellular ‘‘sinks’’ for ATP is attributable to the regula-
tion of ionic fluxes (Hanson and Trewavas, 1982). The role of ATP in mainte-
nance of cell integrity, regulation of cell turgor, phosphorylation of metabolic
substrates, and in other energy utilizing biosyntheses, e.g., the synthesis of
RNA, proteins, membrane lipids, and nucleotide sugars could potentially influ-
ence the rate of fiber growth in many ways (Basra and Malik, 1982).
Adenine nucleotides have scveral effects on respiration. The ratios of different
adenylates is one way of knowing about cellular energy metabolism and its
regulation (Atkinson, 1977). The adenylate energy charge ratio (Atkinson,
1968), (ATP + 0.5 ADP)/(ATP+ADP+AMP), a measure of the energy-rich
adenylates in a cell, was calculated in elongating cotton fibers (Basra and Malik,
1982). It was found that elongating fibers had relatively low energy charge
during the period of rapid cell growth. Generally, growing and dividing cells
maintain a high energy charge around 0.8 whereas senescing or dormant cells
maintain an energy charge of less than 0.5 (Chapman el ul., 1971). In the light of
these data, the critical energy charge threshold for fiber growth seems to be lower
than that of bacterial cells. In spite of the relatively low energy charge values, the
fibers continue to grow at an unarrested rate. It could be emphasized that cell
growth is an energy-consuming system rather than an energy-yielding or storage
one. Therefore, energy charge may be controlled by the rate of ATP usage
whereby low values of energy charge are expected during rapid cell growth.
Pronounced oscillations in individual adenylate ratios during fiber elongation
(Basra and Malik, 1982) reflected that energy charge can obscure large changes
in individual adenylate ratios like ATPIADP, ATPIAMP, and ADPIAMP. Simi-
larly, Lowry et ul. ( I 971) have pointed out that for certain enzymes the ratios
ATPIAMP and ATPIADP may be the metabolically dominant factors rather than
the energy charge per se. The available information on plant metabolism is
insufficient to properly evaluate the role and significance of energy charge
hypothesis.
Higher activities of glycolytic and pentose phosphate pathway enzymes to-
gether with metabolic intermediates and increased rates of turnover of [14C]glu-
cose are observed in elongating fibers of G. hirsutum L. (a long staple type) as
compared to the short staple, G. urboreurn L. (Basra, 1982). Further, pronounced
oscillations along with significant differences in nucleotide ratios suggested rapid
changes in energy metabolic sequences and different metabolic milieu of the two
fibers (Basra and Malik, 1982).The activity of phosphofructokinase and pyruvate
kinase in vivo may be regulated by the rate at which ATP is used (apRees, 1980b;

28. 92 AMARJIT S.BASRA AND C. P.MALIK
Ireland rt ul., 1980). It is pertinent to note that the ATP/ ADP ratio is markedly
lower in the long fibers relative to the short counterpartswhich may be responsible
for the respiratory augumentation ofthe former (Basra and Malik, 1982).Kespira-
tion causes oxidation of substrates to provide energy and the conversion of
substrates to intermediates required for biosynthesis. Therefore, a faster hexose
consumption by long fibers has physiological relevance for attaining increased
length whereas a metabolic depression in terms of hexose oxidation in the short
fibers may dwarf their growth. Overall, nietabolic requirements during fiber
development may be partly met through interrelated operation of the glycolytic
and pentose phosphate pathway.
VIII. Dark Metabolism of Carbon Dioxide
Cotton fiber grows in the dark interior of the boll protected against the en-
vironmental perturbations. Presumably, cotton boll growth results in the produc-
tion of elevated levels of CO, inside the boll. This represents the “external CO,
pool.” As a consequence of growth, an “internal COz pool” would also exist
for each boll constituent, i.e., bur, seed, and lint. The two pools are exchangea-
ble because of free diffusion of the gas across membranes. Several lines of
evidence from various laboratories show that CO, is physiologically a very
active gas, as increased levels of the gas regulate various plant processes (Plumb-
Dhindsa et ul., 1979; Aldasoro and Nicolas, 1980; Bhalla et d.,
1980; Adanis
and Kinne, 1981; Coker and Schubert, 1981; Dhaliwal ct ul., 1981; Ginzburg,
1981; Perez-Tre.jo et al., 1981; Sharma et d.,
1981; Basra and Malik, 1983b).
One of the major effects of increased CO, concentrations is to increase the
rates of CO, fixation which is catalyzed primarily by phosphoenolpyruvate car-
boxylase. Developing cotton fibers possess an active system for assimilating
COz (Dhindsa et a / . , 1975, 1976; Dhindsa, 1978b; Basra and Malik, 1983b).
Studies on unfertilized cotton ovules show that fiber growth is inhibited by the
absence of K + and CO, in the culture environment (Dhindsa et al., 1975). It was
shown that fiber growth is dependent on the turgor pressure in the fiber and that
K + and malate are the osmotically active solutes (osmolytes) which are largely
responsible for the production of turgor in the fiber. Malate is partly synthesized
by dark CO, fixation (Dhindsa et ul., 1975, 1976; Dhindsa, 1978b; Basra and
Malik, 1983b). IAA and GA, affect fiber growth in vitro by regulating the
activities of malate-synthesizing enzymes (Dhindsa, 1978b). ABA, when ap-
plied along with IAA and GA,, inhibits in vitro fiber production by unfertilized
ovules and lowers the malate level in the fibers produced by them (Dhindsa ef
(d., 1976). At least one basis of ABA inhibition of in vitro fiber growth is via
inhibition of malate-synthesizing enzymes by counteraction of GA, effects
(Dhindsa, 1978b).

29. DEVELOPMENT OF THE COTTON FIBER 93
Maintenance of turgor or pressure potential is mandatory for continuous cell
expansion which is achieved by increasing the number of osmolyte molecules in
the cell. Osmoregulation is a process in which turgor is maintained while the
water potential decreases. In elongating cotton fibers, K + and malate levels
fluctuate in correlation with the growth rate and reach peak levels when the
growth rate is maximum (Dhindsa et a/., 1975; Basra and Malik, 1983b). The
parallel behavior of K + and malate during fiber expansion suggested their inter-
relationship as counteracting osmolytes. Maximum concentrations of K + and
malate reached in the fiber can account for over 50% of the osmotic potential of
the fiber (Dhindsa et al., 1975). However, the contribution of soluble sugars,
free amino acids and their derivatives, and other compatible osmolytes to the
osmotic potential of fibers is also important which is as yet undetermined.
Glucose and sucrose are the predominant sugars in fibers and recently “vacuo-
lar” and “cytoplasmic” pools have been observed (Carpita and Delmer, 198I).
Studies on vacuoles from sugarcane suspension cultures show that tonoplast
energization may play a decisive role in both active hexose uptake and active
sucrose uptake at the tonoplast (Thom er a/., 1982; Komor et nl., 1982).There is
a need for knowledge about energetic parameters across the fiber tonoplast as the
fibers are highly vacuolated and solute transfer via the tonoplast and storage in
the vacuole is of great physiological importance in this system.
Photosynthetic cells produce a continual supply of sugars that may influence
their osmotic potential. The lack of photosynthesis in cotton fibers puts the
spotlight on imported and metabolically generated osmolytes, which explains the
large accumulation of K + and malate within the fibers. In a need for carbon
economy, the cotton fibers seem to have successfully exploited the dark metabo-
lism of CO, as an adaptation for intracellular recycling of CO, by refixation to
achieve osmoregulation of growth and to fulfil specific metabolic requirements.
Malate produced by dark fixation pathway possibly acts as an osmoticum and a
counterion for K + accumulation during the period of active fiber extension
(Basra and Malik, 1983b).
The enzymes of malate metabolism, i.e., phosphoenolpyruvate carboxylase
(PEPC), glutamate-oxalacetate-transaminase (GOT), NAD + -malate dehydroge-
nase (MDH), and NADP+-malic enzyme (ME) are readily detected in both in
vitro and in vivo growing fibers (Dhindsa, 197%; Basra and Malik, 1983b).
During the period of rapid growth in vivo, the fibers contain enhanced activities
of PEPC, GOT, and MDH which decline afterward to low values when the rate
of growth slows down (Basra and Malik, 1983b). PEPC directs a portion of the
glycolytic carbon in the form of PEP toward the formation of C, acid, oxalace-
tate. Oxalacetate once formed can either be incorporated into citrate, transami-
nated to aspartate, or reduced to malate. In rapidly elongating fibers, reduction of
oxalacetate to malate is the main route of the metabolism of fixed carbon. This is
evident from the elevated levels of MDH compared with that of GOT, low and

30. 94 AMARJIT S. BASRA AND C. 1’. MALlK
unchanged activity of ME, and quantitative importance of malate during the
period of rapid fiber expansion (Basra and Malik, 1983b). Malate accumulation
in elongating fibers reaches a maximum when the growth rate of fibers is highest
(Dhindsaet d.,
1975;Basra and Malik, 198%). Nevertheless, a marked increase
in GOT concomitant with the period of rapid fiber extension suggested that some
of the oxalacetate formed may serve to provide carbon skeletons for the synthesis
of aspartate (Basra and Malik, 1983b). In addition, oxalacetate and malate pro-
duced by CO, dark metabolism could be consumed as respiratory substrates for
energy-yielding metabolism in fibers. The source of fiber PEP is presumably
glycolysis while the source of CO, is mainly via respiration. The effect of CO,
production may be autocatalytic as one of the metabolic effects of CO, is the
increased utilization of respiratory substrates (Perez-Trejo et ul., I981). The CO,
tension in the cotton bolls has not been determined but is probably high enough
to support high rates of CO, fixation. The carboxylation reaction is endergonic
and hence the advantages to elongating fibers of high rates of CO, fixation need
to be sufficient to outweigh the disadvantages of energy loss.
K + accumulation in elongating fibers has been implied to serve an osmotic
function (Dhindsa et ul., 1975; Leftler and Tubertini, 1976). However, apart
from a role in turgor regulation, changes in K + levels may have more dramatic
cellular consequences. K + is essential as an activator of many enzymes in key
metabolic processes like glycolysis, tricarboxylic acid cycle, oxidative phos-
phorylation, RNA and protein synthesis, etc. (Trewavas, 1976). The regulation
of turgor pressure is related to the vacuolar system. It appears that during the
period of active extension, malate is withdrawn from the metabolic turnover and
is accumulated presumably in the vacuole along with K + as the dication (Basra
and Malik, 1983b).This would maintain the turgor pressure of the fiber as it is
undergoing expansion. Presumably, as the turgor increases with increasing os-
tilotic pressure in the vacuole, the malate housed in vacuole leaks out flooding
the cytoplasm with the substrate. Control of tonoplast influx of other ions by
vacuolar concentration or cell turgor is well establishcd (Cram, 1976).Actually,
during the slowing down of fiber growth, a dramatic increase in ME activity
coupled with a distinct decrease in fiber PEPC activity and malate content is
observed (Basra and Malik, 1983b). ME and PEPC will act antagonistically
being controlled by, among other things, pH and malate concentration (Davies,
1979; Smith and Raven, 1979; Ting, 1981) and these will determine whether
malate is synthesized or broken down. The mechanism generating the driving
force for the reversible accumulation and depletion of malate in the vacuole
remains to be established. A recent attempt to measure transport in vacuoles
isolated fromKulunchiie mesophyll protoplasts has yielded evidence of a specific
malate permease in the tonoplast (Buser-Suter el al., 19x2). This carrier may
merely catalyze the exchange of malate across the tonoplast. ATP and proton
ionophores had no effect on transport rates and hence the malate permease cannot

31. DEVELOPMENT OF THE COTTON FIBER 95
be responsible for malate accumulation. ' Electrophysioh&i~al considerations
have led to the suggestion that malate accumulation in vacuoles is coupled with
the active transport of protons into the cell sap (Liittge and Ball, 1979). It would
be necessary, therefore, to demonstrate the existence of an electrogenically ac-
tive ATPase located in the tonoplast. There is some evidence that malate efflux
from the vacuoles is caused by changes in membrane permeability of the
tonoplast, a decrease in cellular pH, mineral ions like Mg2+, Ca2+,and chang-
ing NADP+/NADPH ratios (Possner er af.,1981).
Increased malate catabolism during later stages of fiber growth is of phys-
iological relevance to meet special requirements of fiber growth. At these stages,
a large proportion of the carbon supply to the fiber is used for the synthesis of a
thick secondary wall and glucose oxidation decreases. MDH and ME together
constitute a transhydrogenase system converting NADH to NADPH. Pyruvate is
produced concomitantly with NADPH by ME reaction and both are important
metabolites for biosynthesis. Therefore, during the later stages of fiber growth
ME reaction may serve to support respiratory activities of the fibers. In this
respect, the dual role of malate as an osmoticurn and a carbon-energy source
during fiber growth seems feasible. Malate is a useful form of stored carbon in
plant cells and can be tolerated at quite high concentrations in the vacuoles.
Amounts of malate synthesizing enzymes, K + , and malate were found to be
higher in elongating fibers of Gossypium hirsutum L. (a long staple type) than the
short counterpart, G. urboreum L. (Basra and Malik, 19831.3).This is supported
by a faster turnover of [14C]glucosein the long fibers resulting in more PEP and
CO, production (Basra, 1982). Therefore, long fibers possess efficient CO,
production and fixation systems relative to short fibers. The capacity for dark
metabolism of CO, may be related to the rate of fiber extension in different
cottons. The ability to generate malate in this way seems to be one important
implication for osmoregulation of growth. It was also observed that during the
period of active elongation, the K + /malate ratio is higher in the arboreumcotton
than the hirsutum cotton (Basra and Malik, 1983b). High K+/malate ratios'of
short fibers are mainly due to the lowered malate levels of these fibers. The
osmolyte systems occur in certain combinations and often in fairly strict propor-
tions. ABA inhibits fiber growth in v i m by inhibiting malate synthesis and
increasing thereby the K+/malate ratio (Dhindsa et ul., 1976). Therefore, the
levels of K+ and malate relative to each other may influence the rate of fiber
growth in the two cultivars (Basra and Malik, 1983b).
It is becoming increasingly appreciated that phosphoenolpyruvate is a major
branch-point in glycolysis (Davies, 1979;apRees, 1980a).Carboxylation of this
intermediate by PEPC is important in CO, and NH, + assimilation to provide a
source of respiratory substrates, amino acids, counterions and NADPH for cell
growth. Each complete turn of the tricarboxylic acid cycle results in the indirect
oxidation of one acetate molecule and the release of two CO, molecules. Thus,

32. 96 AMARJIT S. BASRA AND C. P.MALlK
the number of carbon atoms lost as CO, matches the number of carbon atoms
entering the cycle as acetate, and if this were the only loss the cycle would be
self-sufficient. However, various intermediates are drained from the cycle and
used in biosynthetic reactions of cell growth. Carbon removed in this form must
be replenished for the cycle to continue its operation. Anaplerotic carboxylation
of phosphoenolpyruvateprobably operates and helps to meet this demand. There
seems to be an in vivo competition between PEPC and pyruvate kinase (PK). In
developing soybean seeds, at 27"C, pH 7.5, about 60%of the glycolytic carbon
is processed by PK and 40% by PEPC (Broman et al., 1982).PEPC and PK feed
their products into two separate metabolic pools and it has been suggested that
they should jointly be considered as final enzymes in the glycolytic pathway of
plants (Adams and Rinne, 1981; Broman et al., 1982).The details of the control
of PEP metabolism via PEPC and PK are lacking. The precise fate of the
products of PEPC activity in the dark has yet to be established, but it does seem
clear that such activity must be taken into account in attempts to measure flux
through the pathways of carbohydrate oxidation.
The action of CO, on development is not necessarily by one mechanism alone
but could be achieved by several effects operating in tandem. Some of the CO,
effects may include regulation of the activities of several enzymes (Mitz, 1979;
Arteca and Poovaiah, 1982;Kao and Yang, 1982),regulation of cellular metabo-
lism by changes in pH associated with organic acid content (Davies, 1979),
modification of hormone changes (Dymock and Bown, 1977; Arteca et ul.,
1980; Dhawan et ul., 1981), and possibly other mechanisms.
IX. Cell Walls and Cell Wall Metabolism
The cell walls of higher plants are fundamentally involved in many aspects of
plant biology including the morphology, growth, and development of plant cells.
Primary cell walls are laid down by undifferentiating cells that are still growing
and it is these primary cells that control cell growth. Secondary walls are derived
from primary walls by cells which have stopped or are stopping growth and are
differentiatinginto cells with specialized functions. The cell wall of higher plants
is composed of cellulosic microfibrils embedded in an amorphous matrix of
noncellulosic substances consisting mainly of pectic polysaccharides, hemi-
celluloses, and proteins. During growth of the cell, polymers of the wall interact
and change and the resulting alteration in the properties of the wall can be
correlated with a variation in its function. Since cotton fibers grow synchro-
nously and represent single cells, which are relatively homogeneous with respect
to size, they are especially suitable for studies of the structure and mode of
synthesis of cell walls and of the function of cell walls during growth.
For about one-half of the extension phase, the fiber cell contents are bounded

33. DEVELOPMENT OF THE COTTON FIHt:R 97
by a thin primary wall which undergoes irreversible plastic extension and growth
as the fiber enlarges. Most of the elongation occurs while the fiber is invested
with a primary wall. As the rate of fiber elongation diminishes, the rate of
cellulose deposition at the secondary wall increases. Primary wall eventually
constitutes a thin sheath on the outer surface of the mature fiber. The secondary
wall is deposited on the inner surface of the existing primary wall, i.e., between
primary wall and plasmalemma and at the expense of the cell lumen. The second-
ary wall is less hydrated and more compact than the primary wall and differs
substantially in chemical composition. In a mature fiber, the secondary wall is
about 94% cellulose and relatively thick providing the extraordinary strength
required of a textile fiber (Guthrie, 1955). On both optical microscope evidence
and observations from the electron microscope, the secondary cell wall of ter-
restrial plants is commonly divided into three subdivisions, S, (outermost), S,,
and S,. Normally, the S, layer contains by far the most material. At the present
time, the molecular cause of deposition of three layers and of the orientation of
celluosic microfibrils within them is a matter of some discussion (Robinson,
1977). Primary wall and secondary walls may be distinguished morphologically.
In the former, the cellulosic microfibrils are oriented at a fairly large angle with
respect to the long axis of the cell and form a loosely interwoven network
although some regions exhibit parallel arrangement. The secondary wall is
formed by microfibrils that are more closely packed, lie more parallel to one
another, and are oriented with a smaller angle to the long axis of the cell
(Shafizadeh and McGinnis, 1971).
Cytological, autoradiographic, and biochemical approaches have provided
significant information on the subcellular events of cell wall synthesis. In cotton
fibers, the endoplasmic reticulum, Golgi complex, and plasmalemma form a
functionally integrated membrane system for the synthesis and transport of cell
wall components (Ramsey and Berlin, 1976a,b). Polypeptides of cell wall pro-
teins are synthesized on the rough endoplasmic reticulum, as are other proteins,
however, wall polypeptides are then segregated by release into the lumen of the
endoplasmic reticulum. The polymerization of the hemicellulosic and pectic
polysaccharides from sugar nucleotides and glycosylation of the structural pro-
tein of the wall take place in the dictyosomes after which the products reach the
plasmalemma via vesicles arising from dictyosome cisternae. The vesicles fuse
with the plasmalemma and then open toward the exterior, releasing their contents
to the cell wall by reverse pinocytosis. Westafer and Brown (1976) suggest that
endoplasmic reticulum plays a major role in the synthesis of fiber secondary wall
polysaccharides, and although they report swelling of the endoplasmic reticulum
cisternae and the presence of microvesicles believed to be of endoplasmic re-
ticulum origin, Ryser (1979) could not detect vesicles containing periodate ox-
idizable carbohydrate during secondary wall formation. Again, the main function
of the endoplasmic reticulum might be to deliver to the plasmalemma newly

34. 98 AMAKJIT S.BASRA AND C. P.MALIK
formed polysaccharide synthetases required for secondary wall formation. The
coated vesicles which are often concentrated in the vicinity of the dictyosomes
and also in the cortical cytoplasm just beneath the plasmalemma in plant cells,
occur in developing fibers during the primary and secondary wall formation.
During the secondary wall formation, the number of dictyosome-associated
coated vesicles seems to be smaller than during primary wall formation. Coated
vesicles have been reported from algae to angiosperms (Newcomb, 1979) and
most investigators believe that they are involved in membrane transport.
In the developing fibers, the synthesis of cellulosic microfibrils occurs at the
cell surface, probably in association with multienzyme complexes on the plasma
membrane (Westafer and Brown, 1976; Willison and Brown, 1977). Since the
biochemical isolation of cellulose-synthesizing enzymes has not been accom-
plished, freeze-fracture electron microscopy has been utilized to identify struc-
tures associated with cellulose microfibrils which might play a role in microfibril
synthesis and assembly. In fact, freeze-fracture electron microscopy of the plas-
ma membrane shows close association of certain particles with cell wall micro-
fibrils. The fracture face of the outer leaflet of the plasma membrane shows both
randomly scattered particles and large granules of 250-350 in diameter (Fig.
13)which are correlated with the impressions of microfibrils in the membrane. In
addition, ultrastructural evidence indicates that microtubules near the inner sur-
face of the plasma membrane are arranged parallel to the orientation of wall
microfibrils. Hence the central concept now being advanced is that granular
cellulose-synthesizing complcxes which have lateral mobility in the plane of the
plasma tnembrdne are “spinning” cellulose microfibrils and being guided on
“tracks” composed of niicrotubules.
Recently, terminal ccllulose-synthesizing complexes have been observed on
freeze-fractured plasma membranes of maize and pine seedlings in association
with the ends of microfibril impressions (Mueller and Brown, 1982a). These
terminal complexes are thought to assemblc cellulose microfibrils as they move
through the fluid plane of the membrane (Brown. 1979). The membrane flow
could orient the lateral movement of synthesizing complexes in the membrane
and that microtubules modulate the movement by organizing the microfibrils into
parallel bands in ncwly forming wall layers (Mueller and Brown, 1982b). The
microfilaments made up of actin (a contractile protein) and cortical microtubules
are the structures that are potentially involved in directed microfibril deposition
(Hepler and Palevitz, 1974; Williamson, 1980; Yatsu and Jacks, 1981; Mueller
and Brown, 1982a.b) though the prccise mechanism is not known. Therefore,
more experiments must be done to explore the nature of the cytoskeletal- con-
tractile network and its association with the plasma membrane and to explore
other properties of the plasma membrane which might influence the movement of
terminal synthesizing complexes. The isolation and characterization of terminal
complexes will be one of the most important achievements toward understanding
the microfibril orienting mechanism.

35. DEVELOPMENT OF THE COTTON 1IHliK 99
FIG.13. Thc cxtcrnal fracturc facc (E facc) of thc plasma nicmbranc of developing cotton fibers
as viewed with freeze-fracture electron microscopy. This is how the membrane would appear if
viewed from inside of thc ccll aftcr stripping away thc inncr lcatlct of the plasnia membrane to expose
the outer leaflet closely appressed to the cell wall. Note the impressions of celluloae microfibrils and
of particlc complcxcs (arrows)which arc prcsumcd to function in ccllulosc synthesis. (After Fig. 14,
from Willison and Brown, 1977.)

36. I00 AMARJIT S . BASRA AND C. P. MALIK
It should be pointed out that almost nothing is known about the regulation of
cell wall synthesis. Are the synthesized polymers controlled at the level of
transcription or translation of enzymes, by the levels of substrates, by small
activators or inhibitors. or by hormones'? These questions will remain largely
unanswered until the biosynthetic processes are understood at which time regula-
tion of cell wall biosynthesis will become an exciting area for study of develop-
mental regulation in plants (Darvill et al., 1980).
Long cellulose molecules that make up the walls arejuxtaposed into bundles to
form fibrils (Dolmctsch and Dolmetsch, 1969) which spiral down the length of
the fibers. The microfibrils are held to one another by extensive hydrogen bond-
ing and vary from 10 to 30 nm in breadth depending upon the species. Each
microfibril cross-section contains roughly 10' molecular chains of ( 1 + 4)-p-u-
glucan which exist as extended chains with a twofold screw axis and these are
arranged in an ordcred manner within the microfibril (Preston, 1979). The com-
plex microfibrillar framework in the walls gives the fibers a tensile strength
approximately that of a steel wire of the same diameter. Both the biochemical
and biophysical mechanisms of the assembly of microfibrils are an active area of
study and dispute at the moment (Colvin, 1977, 1981; Burgess, 1979; Franz and
Heiniger, 1981; Robinson, I98I).
The deposition of the fibrils within the walls, i.e., whether thcy are parallel to
the long axis, transverse, random, etc. has an important bearing on the physical
properties of the fibers. Therefore, how the plant synthesizes 6-I ,4-glucan
chains and deposits them in the cell wall is of great interest from an academic and
economic standpoint. Information on the deposition of cellulosic microfibrils in
the fiber walls is scarce. The arrangement of these fibrils in the fibers is very
unusual; rather than exhibiting a continuous spiraling in one direction, they
intermittently reverse their gyre (Betrabet et al., 1963; Roelofsen, 1965). At
these rcversal points, the wall of the fiber is thin and it is here that the mature
fiber twists as it dries and collapses into a ribbon when it is exposed to the air.
The occurrence of twists in the fibers enables them to be spun into yarn. The
mature fiber thus consists of a collapsed tubular cell with a very small lumen and
thick walls composed of superimposed bands of ccllulose, but having thinner
places where the direction of the cellulose spirals was reversed. Microtubules
lining the periphery of the protoplasm tend to parallel the deployment of cell wall
microfibrils and the pattern persists even through the reversals (Yatsu and Jacks,
1981). Colchicine treatment docs not appear to inhibit cellulose synthesis but it
does abolish microtubules in the fiber cells and thus deranges normal wall micro-
fibrillar orientation (Yatsu and Jacks, 198I).
Cell wall synthesis is a developmentally regulated event. The cell wall of the
cotton fiber is a dynamic structure the composition of which is continuously
changing throughout development (Meinert and Delmer, 1977) ending with the
cessation of the fiber's metabolic activity. The primary cell wall is an amalgam

37. DEVELOPMENT OF THE COTTON t:IHtiK 101
of a dynamic equilibrium of component polysaccharides during development but
always has a relatively low cellulose content. The actual amount of cellulose in
primary walls of elongating fibers is a constant value (Beasley, 1979). Cellulose
deposition sharply rises at the onset of secondary wall deposition, declines, and
then rises again (Beasley, 1979). The primary wall is made up largely of pectic
substances and a decrease in the percentage of pectin is recorded with an increase
in the age of the fiber, the rate of decrease being more rapid when the secondary
wall is laid down (Anderson and Kerr, 1938; Whistler eta/., 1940). Changes in
composition of fiber walls during development have been studied from the early
stages of elongation (5 days after anthesis) through the period of secondary wall
formation (16 to 32 days after anthesis) (Meinert and Delmer, 1977). The kinet-
ics of the cell wall was relatively constant until about day 12 when it increased
markedly until secondary wall cellulose deposition was completed. Between
days 12 to 16 after anthesis, increases in essentially all the components contrib-
ute to increase in total wall thickness. At the time of onset of secondary wall
cellulose deposition, a sharp decline in protein and uronosyl residues occurs.
After day 16, deposition of cellulose (and to a much lesser extent increases in
noncellulosic glucose) are the only events which contribute to continuing in-
crease in wall thickening. The deposition of steadily thickening wall takes place
in layers that are alternately denser and lighter under normal day-night condi-
tions, but are absent in fibers developing under constant light (Anderson and
Kerr, 1938) (Fig. 14). The many layers also alternate in the orientation of the
constituent microfibrils and this alternating structure gives tensile strength to
fibers. As in starch formation, diurnal fluctuations in wall deposition might be
expected if photosynthetic sugars were the source of substrate (Fincher and
Stone, 1981).
Huwyler et ul. (1979) found that the absolute amounts of fucose, galactose,
mannose, rhamnose, arabinose, uronic acids, and noncellulosic (3- 1,3-glucan all
reached a maxima at the end of primary wall formation or beginning of second-
ary wall and thereafter decreased, implying that degradation of noncellulosic
polysaccharides was occurring. In contrast, the amounts of xylose and cellulosic
glucan increased until the end of fiber development. A steady decrease in the
percentage of protein content of the walls was observed with increasing age
(Meinert and Delmer, 1977; Huwyler et al., 1979). In a parallel study, Maltby et
al. (1979) found that both soluble and insoluble forms of (3-1,3-glucan increased
in relative amounts up to the time of onset of secondary wall formation. Howev-
er, pulse-chase experiments provided no evidence for subsequent degradation of
the p-1,3-glucan. An acidic arabinogalactan has been isolated from fibers at the
stage of intensive secondary wall formation (Buchala and Meier, 1981).
The characteristics of cellulose biosynthesis in fibers has received extensive
study at the hands of several workers. Marx Figini and her co-workers (Marx
Figini, 1966, 1969; Marx-Figini and Schulz, 1966) showed that in cotton fibers,

38. 102 AMARJIT S. BASRA AND C. P. MALlK
Fic; 14. A transverse section ofa cotton fihcr from a plant grown at 30°C until anlheais and then
sublccted to the regime: light: I hour at JOT: dark: 10hours at 10°C for 1 1 days and then grown for
an additional II days under constant illuniination at B tcmpcraturc of 30°C. After sectioning, the liher
was swollen in cupriethylcnc dianiine. Nurmaaki interference optics. X640. A, Cell lumen; 9,region
of primary wall (not visible). (Photographed by Professor A. B . Wardrop. Department of Botany, La
Trohe Univcrsity, Bundoora, Australia.)

39. DEVELOPMENT OF THE COTTON FIBER 103
the cellulose chains are not only differently oriented in the primary and second-
ary wall, but that they also show different degrees of polymerization. These
workers used viscometric methods to study derivatized glucan chains and sug-
gested that biosynthesis of cellulose consists of two distinct kinetic stages corre-
sponding to the formation of primary and secondary walls during the develop-
ment of the cotton fiber. The first stage proceeds very slowly and yields a small
amount of “primary” cellulose having a nonuniform degree of polymerization
(DPw) ranging from 2000 to 6000. The second stage proceeds much faster and
provides a large amount of “secondary” cellulose having a m w of 14,000.
During the second stage, the m w is independent of variations in the kinetics of
the rate of synthesis of cellulose. The mechanisms by which the degree of
polymerization of the cellulose glucan chains is controlled are unknown.
It is generslly accepted that nucleoside diphosphate sugars (sugar nucleotides)
are the most likely activated monomer substrates for cell wall polysaccharide
synthesis. In cotton fibers, the UDP-sugars are the predominant nucleotide sug-
ars whereas GDP-sugars are not detected in significant quantities (Franz, 1969;
Carpita and Delmer, 1981). The extent of incorporation of UDP-glucose in-
creases with increasing age of the fibers and was found to be very high in fibers
from 20 to 30 days after anthesis (Franz and Meier, 1969). Also GDP-glucose
pyrophosphorylase activity is undetectable but UDP-glucose pyrophosphorylase
activity is found in extremely high levels throughout fiber development (Delmer,
1977). The content of UDP-glucose is maximal at the time of maximum rate of
cellulose deposition in vivo (Carpita and Delmer, 1981). However, it has also
been shown that during the period of rapid elongation and primary wall syn-
thesis, incorporation of radioactivity from GDP-[I4C]glucose into cellulose is
high which gradually declines in older fibers undergoing active deposition of
secondary wall (Delmer et a/., 1974).This has been taken as supportive evidence
that GDP-glucose serves primarily as a precursor for primary wall cellulose. The
question is still open whether two different enzyme systems are involved in
cellulose biosynthesis of primary and secondary walls and how they are reg-
ulated.
The predominant products synthesized from UDP-glucose by extracts of cot-
ton fibers are sucrose, steryl glucosides, p-I ,3-glucan, and cellulose (Delmer et
ul., 1974, 1977; Heiniger and Delmer, 1977) and the rate of synthesis and
turnover of UDP-glucose is more than sufficient to account for the combined
rates of accumulation of these constituents (Carpita and Delmer, 1981). Earlier,
Forsee and Elbein (1972, 1973) demonstrated the synthesis of glucolipids from
UDP-glucose with the biochemical properties of glycosyl phosphoryl-poly-
prenols by using isolated particulate fractions from the cotton fibers taken 12 to
15 days after anthesis. However, the authors were not able to confirm the
intermediate role of the lipid derivatives in the process of cellulose formation.
Conclusive demonstration of the intermediate function of such a compound is

40. I 04 AMAKJIT S. BASRA AND C. P. MALIK
rather difficult, since the known lipid intermediates seem to exhibit a rapid rate
of turnover or low steady-state levels (Carpita and Delmer, 1981). Specific
inhibitors are required to interrupt the reaction sequence at the glycolipid level.
There is now clear evidence in bacterial and mammalian systems that phosphory-
lated prenols are required participants in the synthesis of polysaccharides or
glycoproteins from nucleoside diphosphate sugars (Lennarz and Sher, 1973).
Cotton fibers can also incorporate mannosyl portion of GDP-[14CC]niannoseinto
mannosyl phosphoryl dolichol and lipid linked oligosaccharides (Forsee and
Elbein, 1975). When incubations of the enzymes and GDP-I “Tlmannose are
chased with unlabeled GDP-mannose, the radioactivity is chased from the srnall-
er oligosaccharides into the larger ones suggesting that the smaller oligosac-
charides are precursors for the larger ones (Forsee and Elbein, 1975).The studies
of the synthesis of the mannose containing polymer offer the best evidence in a
plant system for the involvement of a lipid intermediate in the polymerization
reactions. It seems likely that lipid-linked saccharides participate in the synthesis
of glycoproteins, niannan or glucomannan, and cellulose or other glucan syn-
thesis in plants (Ericson and Elbein, 1980). In the chlorophycean alga, Pro-
torheca zop/ii, the newly synthesized ( 1 +
-4) p-glucan chains are transferred
from a lipid-linked intermediate to the protein moiety, suggesting that cellulose
in fact may be synthesized as a glycoprotein (Hopp PI ul., 1978a). Glucolipids
and again oligosaccharide-linked lipids were formed from the substrate UDP-
glucose and endogenous lipids. Indeed, invcstigations on cotton fiber cell wall,
even after thorough purification, show that some protein is attached to the a-
cellulose fraction (Nowak-Ossorio rt d.,
1976;Huwyler rt d . ,1978). However,
the question of possible primers or acceptors for glucan chains remains
unresolved.
Extracts of cell walls from growing cotton fibers contain appreciable amounts
of noncellulosic glucans. Hydrolysis of these glucans by different types of
glucanases and analysis of the degradation products as well as periodate oxida-
tion and methylation studies show that they are essentially composed of p-I ,3-
linked residues (Meinert and Delmer, 1977; Huwyler t>td.,
1978, 1979; Maltby
et cil., 1979). There are no indications of mixed linked p-1JiP- I ,4-glucans
(Huwyler et d.,
1978; Maltby ct d.,
1979). P-3-Iinked glucan synthesis in
cotton fibers is initiated coincident with the earliest stage of secondary wall
cellulose deposition. The first layer of secondary wall has been referred to as the
winding layer in fibers (Kerr, 1946)and it seems likely that most of fl-I ,3-glucan
is associated with this layer (Maltby et al., 1979). As fiber elongation continues
during the period of deposition of the winding layer (Schubert rt ul., 1973, 1976;
Maltby er 01.. 1979) there seem to be differences among cotton varieties in the
extent of elongation which occurs after the onset of secondary wall formation.
Since this period of development coincides with the time of maximal p-I ,3-

41. DEVELOPMENT OF THE COTTON FIHliK I05
glucan deposition, it is possible that the glucans may play some role in determin-
ing the plasticity of the wall at this stage of development.
The synthesis of P-3-linked glucans, commonly referred to as callose, is of
interest in itself. UDP-glucose p-1,3-glucan synthetase has been reported in the
fibers (Delmer et ul., 1977;Heiniger and Delmer, 1977)which is often activated
in vitro by the substrate UDP-glucose and by @-linkeddisaccharides (Delmer,
1977). The early rise of UDP-glucose concentration at the onset of secondary
wall synthesis may be one factor responsible for initiating p-1,3-glucansynthesis
since p-1,3-glucan synthetase is activated by millimolar concentration of UDP-
glucose (Heiniger and Delnier, 1977;Carpita and Delmer, 1981).A recent report
indicates that this enzyme is localized on the plasma membrane of cells of pea
epicotyls (Anderson and Ray, 1978) and preliminary data (reference cited by
Darvill et al., 1980) indicates a similar localization in cotton fibers. Callose is
more commonly found associated with wound responses (Eschrich, 1965)and a
difference in the mode and site of synthesis between wound callose and that
elicited by exogenous UDP-glucose has been observed (Tighe and Heath, 1982).
Callose has been implicated to be an integral but possibly transient component
of fiber cell walls (Meier, 1981). At all stages of secondary wall formation,
callose is synthesized at a very high rate but the total amount in the cell wall does
not exceed 2% in later stages of cell wall formation (Pillonel et al., 1980)due to
a high turnover of this polysaccharide. Subsequently, using intact cotton fibers,
Meier et al. (1981) have provided corroborative evidence for a clear turnover of
callose. These workers suggested callose to be an intermediate in cellulose
synthesis at the stage of secondary wall formation and it was implied that callose
may accumulate where the biosynthetic step from callose to cellulose is inhib-
ited. However, the results do not show whether callose is a direct glucosyl donor
for cellulose. If this is the case, a (1 + 3)-P-~-glucan:(l-4)-p-~-glucan
glucosyltransferase (transglucosylase) would probably be involved in cellulose
synthesis. Waterkeyn (1981) has shown by fluorescence microscopy, after stain-
ing with aniline blue, that callose is always localized independently of the fiber
age in the innermost wall layer bordering the cell lumen from the onset of the
secondary thickening until the end of fiber development. It was proposed that
callose may be performing the precursor role to cellulose biosynthesis or, more
probably, as forming a permanently restored interface or matrix where cellulose
microfibrils undergo a sort of maturation and are oriented before their incorpora-
tion in the organized cell wall. However, little is known about the in vivo
regulation whereby callose synthesis is initiated coincident with the onset of
secondary wall synthesis. At the early stages of secondary cell wall formation,
glucose and fructose exhibit a maxima which was closely followed by a maxima
in the (1 .--, 3)-P-~-glucan
content and in the sugar phosphates (glucose 1-
phosphate, glucose 6-phosphate, fructose 6-phosphate) (Jaquet et al., 1982).