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PROPAGATION OF PECAN (CARYA
ILLINOENSIS) USING IN VITRO
TECHNIQUES
ADEELA HAROON
DEPARTMENT OF BOTANY UNIVERSITY OF THE PUNJAB
LAHORE, PAKISTAN
PROPAGATION OF PECAN (CARYA
ILLINOENSIS) USING IN VITRO TECHNIQUES
A THESIS SUBMITTED TO THE UNIVERSITY OF THE PUNJAB
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY IN
BOTANY
BY
ADEELA HAROON
DEPARTMENT OF BOTANY
UNIVERSITY OF THE PUNJAB LAHORE, PAKISTAN
JULY, 2010
DEDICATED TO:
My Father Hon/Capt. (R) Haroon-ur-Rashid
TK (MIL)-Cl-I
Who is every thing,
My consolation in sorrow,
My hope in misery,
My strength in weakness,
That only one,
who sacrificed his life for my effluent future.
My Mother Salma Haroon
Whose inspiration towards knowledge served me,
as a Beacon of Light whom I own all,
That is mine.
My Brother Abdul Moueed Haroon
My Sisters Munazah, Fareeha, Narmeen, Wajeeha Noor
for their subtime love and deep affection.
CONTENTS
TITLE PAGE NUMBER
CERTIFICATE i
ACKNOWLEDGEMENTS ii
ABBREVIATIONS/ UNIT ABBREVIATIONS iii
ABSTRACT v
LIST OF ANNEXURE viii
LIST OF FIGURES ix
LIST OF TABLES xxiv
Chapter 1: INTRODUCTION 1
Chapter 2: LITERATURE REVIEW 5
2.1 Tissue Culture Studies in Pecan 6
2.1.1 Micropropagation 7
2.1.2 Somatic Embryogenesis 9
2.1.3 Novel Micropropagation Methods 17
2.1.4 Adventitious Regeneration 23
2.1.5 Effect of TDZ 26
Chapter 3: MATERIALS AND METHODS 29
3.1 Media Preparation 29
3.1.1 Preparation and Storage of Stock Solutions 29
3.1.2 Growth Regulators 30
3.1.3 Preparation of Stock Solutions for DKW (Driver and Kuniyuki,
1984 Medium) 30
3.1.4 Preparation of Stock Solutions for MS (Murashige and Skoog,
1962) Medium 32
3.1.5 Preparation of Stock Solutions for WPM (Woody Plant Medium
Of McCown and Llyod, 1981) 33
3.1.6 Preparation of Stock Solutions of Growth Regulators 35
3.1.7 Preparation of DKW Medium from the Stocks 35
3.1.8 Preparation of MS and WPM Medium from the Stocks 35
3.2 Sterilization 36
3.2.1 Glassware Sterilization 36
3.2.2 Sterilization of Tissue Culture Media 37
3.2.3 Sterilization of Glasshouse Media 37
3.2.4 Sterilization of Working Area of Laminar Airflow Cabinet 37
3.2.5 Sterilization of Surgical Tools 38
3.3 Plant Material 38
3.3.1 Source of Plant Material 38
3.3.2 Disinfestation of Plant Material 38
3.4 Experimental Plan 39
3.4.1 In vitro Germination of Pecan Seeds 39
3.4.2 Callus Induction and Organogenesis 42
3.4.2.1 Callus Induction from Bark Segments 42
3.4.2.2 Callus Induction from Immature Fruits 43
3.4.3 Adventitious Regeneration from Immature Cotyledonary
Explants of Pecan 44
3.4.4 Novel Micropropagation Protocols 44
3.4.4.1 Shoot Forcing 44
3.4.4.2 Forcing Large Stem Segments 45
3.4.4.2.1 Establishment of Softwood Shoots In
Different Rooting Medium 46
3.4.5 Inoculation of Explants 46
3.4.6 Culture Conditions 47
3.4.7 Statistical Data Analysis 47
Chapter 4: PRODUCTION OF PECAN SEEDLINGS FOLLOWING
IN VITRO GERMINATION 48
RESULTS 48
4.1 In vitro Germination of Pecan Seeds 48
4.2 Rooting of In vitro Multiple Shoots Developed from In vitro-
Grown Seeds of Pecan 58
4.3 Hardening and Acclimatization of In vitro-Grown Plants of Pecan 62
DISCUSSION 66
Chapter 5: NOVEL MACRO/ MICROPROPAGATION METHODS 73
RESULTS 73
5.1 Shoot Forcing 73
5.2 Forcing Large Stem Segments 77
5.2.1 Establishment of Softwood Shoots in Different
Rooting Medium 98
DISCUSSION 101
Chapter 6: CALLUS INDUCTION AND ORGANOGENESIS 110
RESULTS 110
6.1 Callus Induction from Bark Segments 110
6.2 Callus Induction from Immature Fruit 119
DISCUSSION 131
Chapter 7: ADVENTITIOUS REGENERATION OF PECAN USING IMMATURE COTYLEDONARY EXPLANTS 137
RESULTS 137
DISCUSSION 143
Chapter 8: GENERAL DISCUSSION AND FUTURE WORK 147
Chapter 9: LITERATURE CITED 153
ANNEXURES (I-XII) 180
ANNEXURE (Published Article- I) 188
i
CERTIFICATE
This is to certify that the research work entitled “Propagation of Pecan (Carya
illinoensis)” using in vitro techniques” described in this thesis by Ms. Adeela Haroon is
an original work of the author and has been carried out under my direct supervision. I
have personally gone through all the data, results, materials reported in the manuscript
and certify their correctness and authenticity. I further certify that the material included
in this thesis has not been used in part or full in a manuscript already submitted or in the
process of submission in partial or complete fulfillment of the award of any other degree
from any institution. I also certify that the thesis has been prepared under my supervision
according to the prescribed format and I endorse its evaluation for the award of Ph.D
degree through the official procedures of the University of the Punjab, Lahore.
ii
ACKNOWLEDGEMENTS
He, who dares, seeks knowledge.
All praises and hymns are for ALLAH, the lord of creation, the compassionate,
the merciful, the ruler of the day of judgment, the creator and cherisher of the world. He
is the one, who guided through my research work and revealed to me secrets of nature.
I would like to record my sentiments of indebtedness to my respected supervisor,
Dr. Faheem Aftab, Associate Professor, Department of Botany, University of the
Punjab, Lahore who guided with gratitude and sympathetic behavior throughout my
research work. His scholarly guidance, constructive criticism, dedicated interest, co-
operation and encouragement was the real source of motivation for me during my
research work.
It is honour for me to express my deep indebtedness to Prof. Dr. Rass Masood
Khan, Chairman, Department of Botany, University of the Punjab, Lahore for providing
best research facilities to accomplish this research work.
I have no words to explain my heartiest feelings to Dr. Humera Afrasiab
(Assistant Professor, Botany Department) who encouraged throughout my research work
and also directed my conscience in a real direction.
I must acknowledge my heartily thanks to my lab fellows Dr. Neelma Munir,
Zahoor Ahmad Sajid, M. Akram for their vital instructions and intellectual suggestions
during the course of my research and in the submission of this thesis.
My sincere thanks are to Mrs. Sadia Rizwan and Ms. Arifa Khalid who
willingly helped me to achieve this goal.
My deep sentiments to my father for his spiritual and financial support during my
research work.
I feel fully justified to pay my regards to my friends Dr. Khajista Jabeen,
Naveeda Batool for their perpetual help through the completion of my thesis.
ADEELA HAROON
iii
ABBREVIATIONS/ UNIT ABBREVIATIONS
µM Micromolar
µmol m-2 s-1 Micromole per meter square per second (Light Intensity)
2, 4-D 2, 4-Dichlorophenoxy acetic acid
8-HQC 8- Hydroxyquinoline Citrate
ABA Abscisic Acid
AFLP Amplified Fragment Length Polymorphism
ANOVA Analysis Of Variance
BA/ BAP 6-Benzyladenine/ aminopurine
BDS Basal Dunstan and Short, (1977) Medium
ºC Degree Centigrade
cm Centimeter
Conc. Concentration
cv. Cultivar
cvs. Cultivars
df Degrees of Freedom
DKW Driver and Kuniyuki (1984) Walnut Medium
DMSO Dimethyl sulfoxide
EDTA Ethylenediaminetetraacetic acid
Fig. Figure
GA3 Gibberellic Acid
g l-1 Gram per liter
g Gram
h Hour
HCl Hydrochloric Acid
HgCl2 Mercuric chloride
H2O2 Hydrogen peroxide
H2SO4 Sulphuric acid
IAA Indole-3-acetic acid
IBA Indole-3-butyric acid
K2Cr2O7 Potassium dichromate
KOH Potassium Hydroxide
l Liter
iv
L. Linnaeus
lbs inch-2 Pounds Per Square Inch
LS Linsmaier and Skoog, 1965 Medium
M Molar
mg Milligram
mg l-1 Milligram per liter
ml Milliliter
ml l-1 Milliliter per liter
mM Millimolar
mM l-1 Millimoles per liter
mm Millimeter
MS Murashige and Skoog (1962) basal medium
N Normal
NAA Naphthalene acetic acid
NaOCl Sodium hypochlorite
NaOH Sodium Hydroxide
nM Nanomolar
N.W.F.P North-West Frontier Province
PGRs Plant Growth Regulators
pH Power of Hydrogen ion concentration
ppm Parts per million
RAPD Random Amplified Polymorphic DNA
SE Standard Error
spp. Species
SPSS Statistical Package for Social Sciences
SSR Simple Sequence Repeat
TDZ Thidiazuron (N-phenyl-N'-1, 2, 3-thidiazol-5-yl-urea)
UV Ultraviolet
v/v Volume/Volume
w/v Weight/Volume
Wan. Wangenheim
WPM Woody Plant Medium of McCown and Llyod (1981)
v
ABSTRACT
During the present work, an in vitro approach was followed for the propagation
of Pecan [Carya illinoensis (Wangenheim) C. Koch]. Effect of different media (DKW,
MS or WPM) supplemented with various levels of BAP were tested for in vitro
germination response. It was observed that the best medium for in vitro germination was
an MS formulation supplemented with 4.0 µM BAP. During the experiments, different
morphological features of in vitro-grown seedlings of Pecan were also observed.
Formation of multiple shoots was also observed from intact nodal regions of developing
seedlings. Multiple shoots developed from in vitro germinated seeds were shifted to
various rooting media. After acquiring a sufficient length (3 - 4 cm), the developed
multiple shoots were transferred to the rooting media, i.e., DKW or MS supplemented
with different combinations of growth regulator (IAA, IBA or NAA). MS medium
supplemented with 4.0 µM IBA + 4.0 µM NAA proved to be best medium for root
induction. On the other hand, in vitro-germinated seedlings after acquiring a sufficient
length (4 - 5 cm), were transferred successfully to perlite or vermiculite to enhance
rooting. More than 85 % of in vitro-grown Pecan plants were acclimatized successfully
to the soil under glasshouse conditions and kept for more than 30 days. These plants
were then transferred to field conditions at Botanical Garden, Punjab University, Lahore.
For the clonal propagation of Pecan, forcing shoot tips and/ or epicormic buds
from the large stem segments taken from more juvenile portions of older trees. Softwood
shoots were forced form shoot tips of Pecan during the dormant season. Forcing solution
(8-HQC) containing sucrose (2 %), TDZ (2.0 µM) in combination with IBA (2.0 µM)
and BAP (2.0 µM) was quite effective for the highest (89.45 %) sprouting of buds under
glasshouse conditions. Softwood shoots were also forced through epicormic or latent
buds form the large branch segments on different media under various environmental
vi
conditions. The present investigation demonstrated that glasshouse conditions favored
the maximum (2.92) production of softwood shoots as compared to other lab or wire
house (natural) conditions. Media also had a significant effect on softwood shoot
production as sterilized sand produced the highest (2.92) mean number of softwood
shoots during the winter season. Rooting experiments with these softwood shoots
however were not successful as contamination was the major limitation.
During the present investigation, a suitable explant for callus induction and its
subsequent maintenance was established for Pecan (Carya illinoensis Wan.). Bark
segments and immature cotyledons of Pecan were used as an explant source. Effect of
different media (DKW, MS or WPM) supplemented with various levels of 2, 4-D, NAA
and TDZ were tested for callogenesis from bark and cotyledonary explants. Mature bark
explants cultured on DKW medium containing a combination of 2, 4-D and TDZ
resulted in 93.70 % callus induction and proved to be the best medium for callus
induction and its maintenance. DKW medium supplemented with 13.57 µM TDZ
resulted in 93.33 % callus induction from immature fruit explants. Morphologically
different calluses were also observed at various levels of 2, 4-D, NAA and TDZ. Tissue
browning was a major problem associated with callus induction using bark and fruit
explants. These brown callus cultures, however, formed root primordia during this study
after an incubation period of 110 days or so. Calluses were also transferred for plant
regeneration, however, there were no plantlet regeneration possible during the present
investigation. In the present study, adventitious multiple shoots were initiated from
immature cotyledonary explants of Pecan (Carya illinoensis). The embryo axes were
excised carefully and small cotyledonary pieces from immature fruits were cultured on
different media (DKW, WPM or MS) supplemented with various levels of benzyl
aminopurine (BAP) or Thidiazuron (TDZ). The shoots were initiated after 8 days from
vii
the cultures on DKW or MS media supplemented with BAP (0.5, 1.0, 4.0, 8.0 or 15 µM).
Media supplemented with TDZ had shown no response. Maximum shoot development
and proliferation was observed after 16 days of culture on MS medium supplemented
with 15 µM BAP. The developed shoots were then transferred to basal MS medium for
further development and shoot proliferation for 20 days. The proliferated shoots (2.0 -
4.5 cm long) were transferred (without any pre-treatment) to fresh MS basal medium for
root initiation. Although rooting could not be achieved during this research work, some
progress has been made in this regard. However, adventitious regeneration indicates a
strong possibility to regenerate whole plants from various tissues of Pecan.
viii
LIST OF ANNEXURES
ANNEXURE TITLE PAGE
NUMBER NUMBER I: Formulation of DKW Medium (Driver and Kuniyuki, 1984)
for the preparation of stock solutions 180
II: Formulation of MS Medium (Murashige and Skoog, 1962)
for the preparation of stock solutions 181
III: Formulation of WPM Medium (Woody Plant Medium of
McCown and Llyod, 1984) for the preparation of stock
solutions 182
IV: Growth regulators used in the study with respective
abbreviation, molecular weight and initial solvent 183
V: Preparation of 1 liter DKW Medium 184
VI: Preparation of 1 liter MS Medium 184
VII: Preparation of 1 liter WPM Medium 185
VIII: Composition of different media used for in vitro germination
of Pecan seeds 185
IX: Composition of different media used for callus induction/
maintenance from mature bark 186
X: Composition of different media used for plant regeneration
from callus cultures 186
XI: Composition of different media used for callus induction/
maintenance from immature fruits 187
XII: Composition of different media used for adventitious
regeneration of Pecan 187
ix
LIST OF FIGURES
FIGURE TITLE PAGE
NUMBER NUMBER
3.1: A mature fruit of Pecan placed on a clean glazed ceramic
slab (0.5 x). 39
3.2: An enlarged view of Pecan fruit after removing the husk (1.0 x). 39
3.3: Excised one third fruit of Pecan showing the excision of right
cotyledon longitudinally parallel to the embryonal axis (0.6 x). 39
3.4: A view of fruit excised parallel to the right side of embryonal
axis showing the removed cotyledon (arrow) (0.6 x). 40
3.5: Another view of fruit excised parallel to the left side of
embryonal axis showing the removed cotyledon (arrow)(0.6 x). 40
3.6: An enlarged longitudinal view of Pecan fruit excised on both
sides parallel to the embryonal axis (0.8 x). 40
3.7: A view of excised Pecan fruit showing the removal of brown
testa of fruit from the upper part of the embryonal axis (0.6 x). 40
3.8: An enlarged view after the removal of brown testa on the upper
part of the embryonal axis highlighting the exposure of embryo
(arrow) (0.7 x). 40
3.9: A culture vessel showing the embryonal axis with the one third
fruit’s cotyledonary portion inoculated in respective medium
(0.5 x). 40
4.1: Root initiation (arrow) on WPM medium at day 6 of initial
culture, showing also the cotyledonary portion of cultured fruit
(right bracket) (1.6 x). 52
4.2: Elongation of root (in the direction of curved arrow) on WPM
medium at day 8 of initial culture (1.3 x). 52
4.3: Simultaneous development of shoot and root on WPM medium
supplemented with 1µM BAP at day 6 of initial culture (1.3 x). 52
4.4: Elongation of root (curved arrow) on DKW medium at day 8 of
initial culture (1.0 x). 52
x
4.5: Elongation and further development of root at day 9 of initial
culture. An arc shows the cotyledonary portion of fruit cultured
on DKW medium (1.3 x). 52
4.6: Another view (almost at right angel to the view in Fig. 4.5)
showing the development of root (2.0 x). 52
4.7: Simultaneous development of both shoot and root (double
headed arrow) on MS basal medium at day 9 (1.3 x). 53
4.8: Development of shoot (shorter arrow) and root (violet arrow)
with the formation of callus (a curved line over green area) from
the fruit portion adjoining the embryonal axis on DKW medium
supplemented with 4 µM BAP at day 15 (Front view, 1.3 x). 53
4.9: An enlarged view of Fig. 4.8 highlighting the formation of callus
(arrows) from the fruit portions adjoining the embryonal axes on
DKW medium supplemented with 4µM BAP at day 15 (2.5 x). 53
4.10: An opposite view of Fig. 4.8 showing the development of root
(arrow) at 1.3 x. 53
4.11: Multiple shoot formation (curve) on DKW medium
supplemented with 4 µM B (0.5) x). 55
4.12: An enlarged and opposite view of multiple shoots (brace)
formed on DKW medium supplemented with 4µM BAP. An
arrow indicates the remaining fruit portion (1.3 x). 55
4.13: Multiple shoot formation (arrows) on DKW medium
supplemented with 8 µM BAP (0.7 x). 55
4.14: Multiple shoot (small arrows) formation on WPM medium
supplemented with 4 µM BAP, a bigger arrow indicating the
direction and development of root (1.0 x). 55
4.15: An opposite view of Fig. 4.14 (curved arrow) indicating the
direction of further development of root from in vitro
germinating seedling of Pecan (1.0 x). 55
4.16: A left side view of Fig. 4.15 showing the vigorous root
development (a longer arrow). A short arrow indicates
secondary root development on WPM medium supplemented
with 4 µM BAP (1.0 x). 55
xi
4.17: A bunch of multiple shoots arising from single initial point on
DKW medium supplemented with 8 µM BAP also showing
the remaining fruit portion (0.9 x). 56
4.18: Two shoots (double headed arrow) developed on MS medium
supplemented with 12 µM BAP, also showing the cotyledonary
portion of fruit (two single arrows) (1.3 x). 56
4.19: In vitro germinating Pecan seedling showing maximum shoot
length with upward root highlighting two nodular structures
(arrow) at the root tip formed on WPM medium supplemented
with 12 µM BAP (0.7 x). 56
4.20: Development of primary root with the formation of secondary
roots (right bracket, arrows) on DKW medium supplemented
with 4 µM BAP at day 25 (front view to the embryonal axis)
(0.3 x). 57
4.21: An opposite and full view of Fig. 4.20 showing the formation
of multiple shoots (arrows) and nodular Structures (left bracket)
on secondary roots at day 25 of initial culture (0.5 x). 57
4.22: An enlarged view of Fig. 4.21 (lower half) showing the
formation of nodular structures (arrows) on secondary roots
at day 25 of initial culture (1.0 x). 57
4.23: Multiple shoots transferred to MS medium supplemented
with 8 µM NAA showing the formation of light-green friable
callus with white patches at the shoot base(arrow) at day 15
of transfer to rooting medium (1.0 x). 59
4.24: Shoot transferred to DKW medium supplemented with
2 µM IBA showing the formation of friable, transparent and
brown callus at the shoot base (arrows) at day15 of transfer
to rooting medium (1.0 x). 59
4.25: Shoot transferred to MS medium supplemented with 4 µM
IBA + 4 µM NAA showing the formation of compact,
yellowish-brown callus at the shoot base and developed root
(arrow) after 35 days of transfer to rooting medium (0.8 x). 59
xii
4.26: Formation of two roots (arrows) with brownish, compact
callus at the shoot base on MS medium supplemented with
4 µM NAA at day 32 of transfer to rooting medium (0.9 x). 59
4.27: An opposite view of Fig. 4.26 highlighting the formation of
callus, roots and multiple shoots (single arrows) (0.9 x). 59
4.28: Formation of two roots (two combined arrows) with
yellowish-brown, compact callus at shoot base on MS
medium supplemented with 4 µM IBA + 4 µM NAA at day
30 of transfer to rooting medium (1.0 x). 59
4.29: Formation of compact, transparent, watery callus (arrows) at
shoot base with no root on MS medium supplemented with 4
µM IBA + 4 µM NAA at day 26 of transfer to rooting
medium (1.0 x). 59
4.30: An enlarged and opposite view of Fig. 4.29 showing the
formation of creamy-white callus (arrows) at base of multiple
shoots (MS-arrows) (1.3 x). 59
4.31: Root induction (arrow) with transparent, yellowish-brown,
compact callus formed (double arrows) on MS medium
supplemented with 4 µM IBA + 4 µM NAA at day 25 (1.2 x). 59
4.32: Vigorous callus growth with development of root (longer arrow)
on MS medium supplemented with 4 µM IBA + 4 µM NAA at
day 32 showing shoot necrosis and ultimately shoot death
(shorter arrow) (0.9 x). 60
4.33: Browning of callus, swelling and browning of the root at day
37 of transfer to rooting medium (0.9 x). 60
4.34: Multiple shoots with the formation of compact, brown callus
showing root initiation (arrow) on MS medium supplemented
with 4 µM NAA at day 25 of transfer to rooting medium (0.7 x). 60
4.35: An enlarged view of Fig. 4.34 highlighting the root induction
(arrow) and necrosis of shoots showing no further growth of
root at day 35 (0.9 x). 60
4.36: In vitro-grown Pecan seedling transferred in perlite showing
the remaining cotyledonary part (arrow) of fruit at day 1 (0.5 x). 62
4.37: The Pecan plantlet in perlite at 7th day (0.3 x). 62
xiii
4.38: In vitro-grown Pecan seedling being hardened in vermiculite
showing the remaining cotyledonary part (arrow) of fruit at
day 1 (0.8 x). 63
4.39: The Pecan seedling at 7th day of transfer in vermiculite (0.3 x). 63
4.40: The in vitro-grown Pecan seedlings in perlite and vermiculite
kept in an artificially constructed chamber (arrow showing
polyethylene sheet) for retention of humidity (0.1 x). 63
4.41: Browning of the leaves has just begun from the tip (arrows)
in vermiculite at day 11 (1.0 x). 64
4.42: Browning of the leaves extended towards the leaf base in
vermiculite at day 19 (1.3 x). 64
4.43: The death of Pecan plantlet at day 27 (1.3 x). 64
4.44: In vitro-grown Pecan plantlets in perlite and vermiculite
kept in an artificially constructed chamber (polyethylene
sheet) at day 15 (0.2 x). 64
4.45: In vitro-grown Pecan plantlets at day 30 (0.2 x). 64
4.46: Pecan plant kept in culture room for 15 days at
25 ± 2 °C after acclimatization (0.3 x). 65
4.47: Acclimatized Pecan plant under glasshouse conditions
at 45th day under natural light conditions 25 ± 2 °C (0.25 x). 65
4.48: Acclimatized Pecan plant in glasshouse at 65th day (0.25 x). 65
4.49: Acclimatized well developed in vitro-raised plants of Pecan
ready for their transfer to field conditions (0.15 x). 65
5.1: A mature Pecan tree from Lahore, Pakistan. A) photographed
in May, 2008. B) the same tree as in A during dormant season
(December, 2008). 74
5.2: Effect of different environments and media formulations on
shoot forcing from Pecan stems segments (25 cm long) during
the spring season (February - March). 74
5.3: Pecan shoot segments immersed in forcing solution, i.e.,
distilled water with 200 mg/l 8-HQC, 30 g sucrose and IBA +
TDZ both at a concentration of 0.5 µM and BAP (5.0 µM)
under culture room conditions (1.0 x). 75
5.4: An enlarged view of the marked part from Fig. 5.3 (1.0 x). 75
xiv
5.5: Shoot segments immersed in distilled water with 200 mg/l
8-HQC, 30 g sucrose, IBA + TDZ (2.0 + 2.0 µM) and
BAP (15.0 µM) under culture room conditions (1.0 x). 75
5.6: An enlarged view of dotted central part from Fig. 5.5 (1.3 x). 75
5.7: Pecan shoots immersed in glass-jars highlighting the
swelling of buds (arrows) in forcing solution containing
IBA + TDZ (0.5 + 0.5 µM) and BAP (5.0 µM) under
wire house conditions at day 6 (1.0 x). 76
5.8: Pecan shoots immersed in glass-jars indicating the swelling
of buds appearing green in colour (arrows) in medium
containing IBA + TDZ (0.5 + 0.5 µM) and BAP (5.0 µM)
in glasshouse at day 6 (1.0 x). 76
5.9: Dotted area highlighting swelling of buds appearing bright
green from shoots placed in forcing solution containing IBA
+ TDZ (2.0 + 2.0 µM) and BAP at 15.0 µM under glasshouse
conditions at day 6 (1.0 x). 76
5.10: An enlarged view of highlighted area from Fig. 5.9 (2.5 x). 76
5.11: Initiation of sprouting (arrows) of shoots in logs placed on
sterilized sand, a broader arrow indicating the enlarged
view of the sprouted buds at day 9 (1.0 x). 79
5.12: Further development of sprouted epicormic buds seen
in Fig. 5.11 in to male inflorescence, “catkin” (arrows)
at day 17 (1.0 x). 79
5.13: An enlarged view of log placed in sand showing sprouting
of multiple buds (1.0x). 79
5.14: Multiple sprouting observed in logs placed on sterilized
coccopeat, arrow pointing towards enlarged view (1.0 x). 79
5.15: Development of multiple shoots (arrows) in logs on
sterilized coccopeat (1.0 x). 80
5.16: Emergence of multiple shoots (arrows) in logs on sterilized
sawdust (1.0 x). 80
5.17: Pecan logs placed in flats filled with sterilized coccopeat
showing the sprouting of buds and development of
softwood shoot (highlighted areas) (1.0 x). 80
xv
5.18: Enlarged views of the highlighted areas from Fig. 5.17. 80
5.19: Sprouting of epicormic buds observed in sawdust (arrows)
at day 8 (1.0 x). 81
5.20: Pecan logs placed in flats filled with sterilized sand showing
the sprouting of buds (arrows), at day 8 at 25 ± 2 ºC (1.0 x). 81
5.21: Sprouting of epicormic buds (arrows) in coccopeat at day 8
at 25 ± 2 ºC (1.0 x). 81
5.22: Sprouting of epicormic buds (black arrows) and softwood
shoot (brown arrows) development from logs placed in
sterilized sand at day 21 (1.0 x). 82
5.23: Sprouting of buds (arrows) in logs placed in sterilized sawdust
at day 8 (1.0 x). 82
5.24: Pecan logs placed in flats filled with sterilized coccopeat.
Forcing epicormic buds using this medium was possible as
shown here at day 8 (1.0 x). 82
5.25: Sprouted epicormic buds (brown arrows) and softwood
shoots (black arrows) developed in sand at day 11 (1.0 x). 84
5.26: Sprouting and growth of soft-wood shoots from logs placed in
sand. A & B) A view of flat filled with sterilized sand (1.0 x).
C) An enlarged view of the highlighted area from A (1.2 x).
D) An enlarged view of the highlighted area from B (2.1 x). 86
5.27: Forcing of epicormic buds and development of softwood
shoots in logs placed in flats filled with sterilized sand at
day 47 (1.0 x). 87
5.28: A softwood shoot from logs placed in sterilized sawdust at
day 47 (1.0 x). 87
5.29: Softwood shoots (arrows) from logs placed in flats filled
with sterilized coccopeat at day 47 (1.0 x). 87
5.30: Pecan logs placed in flats filled with sterilized sand,
coccopeat and sawdust showing the sprouting and growth
of softwood shoot at day 51 (1.0 x). 88
5.31: Effect of different seasons on epicormic bud induction potential
with reference to bud-derived shoot parameters in Pecan logs.
Vertical bars above the columns are the SE (±)of the means.
xvi
Different letters above the vertical bars representing the
significant differences according to Duncan’s Multiple Range
test at P<0.05 value. 90
5.32: Effect of different media on epicormic bud induction potential
with reference to bud-derived shoot parameters in Pecan logs.
Vertical bars above the columns are the SE (±) of the means.
Different letters above the vertical bars representing the
significant differences according to Duncan’s Multiple Range
test at P<0.05 value. 91
5.33: Effect of different environments on epicormic bud induction
potential with reference to bud-derived shoot parameters in
Pecan logs. Vertical bars above the columns are the SE (±) of
the means. Different letters above the vertical bars representing
the significant differences according to Duncan’s Multiple
Range test at P<0.05 value. 92
5.34: A cumulative effect of media, environment and season on
forcing potential of the logs regarding the parameters studied
(number of sprouts, shoots, nodes, leaves and shoot length) in
Pecan. Vertical bars above the columns are the SE (±) of the
means. Different letters above the vertical bars representing
the significant differences among different values according
to Duncan’s Multiple Range test at P<0.05 value. This figure
depicts the cumulative data of three experiments. In each
experiment 9 logs were placed in three media (each medium
has 3 trays and 3 logs per tray) under three environmental
conditions during three seasons. 93
5.35: Pecan logs placed in sand indicating the formation of callus
(dotted area) at the cut surfaces under glasshouse conditions
at day 17 (1.0 x). 94
5.36: An enlarged view of the dotted portion from Fig. 5.35 (1.6 x). 94
5.37: Another photograph showing the formation of callus (arrows)
at the cut surface of log placed in sand under glasshouse
conditions (1.0 x). 94
xvii
5.38: Pecan log indicating the formation of callus (arrows) at the
cutting points in coccopeat under glasshouse conditions. 95
5.39: An enlarged view of the highlighted portion from Fig. 5.38
(1.0 x). 95
5.40: Pecan logs placed in sand indicating the formation of callus (an arc)
at the cut surfaces in sand under culture room conditions (1.0 x). 95
5.41: An enlarged view of the highlighted portion from Fig. 5.40
(1.0 x). 95
5.42 - 5.43: Pecan logs placed in sawdust, dotted areas indicating the
presence of contamination on the media surfaces under culture
room conditions (1.0 x). 96
5.44: Pecan logs placed in sawdust, arrows indicating the contamination
of sprouted buds under glasshouse conditions (1.0 x). 97
5.45: Pecan logs placed in sterilized sand, the highlighted dotted
areas indicating the presence of contamination on the media
surfaces under culture room conditions (1.0 x). 97
5.46 - 5.47: Soft wood shoots derived from epicormic/ latent buds placed
in sterilized sand for rooting phenomenon under culture room
environment at 25 ± 2 ºC (1.0 x). 98
5.48: A soft wood shoot harvested from forced Pecan logs placed
in peat moss for rooting under controlled environmental
conditions (1.0 x). 99
5.49-50: Soft wood shoots harvested from forced Pecan logs placed
in different grades of vermiculite for rooting under controlled
environmental conditions (1.0 x). 99
5.51: A soft wood shoot harvested from forced logs with the leaves
removed placed in vermiculite for rooting under controlled
environmental conditions (1.0 x). 99
5.52: A comparison of different rooting media with softwood
shoots maintained in culture room at 25 ± 2 ºC (1.0 x). 99
5.53: Plastic pots containing softwood shoots were placed under an
artificially constructed chamber with transparent polyethylene
sheet for the maintenance of high humidity kept in culture
room at 25 ± 2 ºC (1.0 x). 100
xviii
5.54: A plastic pot showing fungal contamination (arrow) along
the base of dried softwood shoot kept in culture room at
25 ± 2 ºC (1.0 x). 100
5.55: A plastic pot showing fungal contamination spread on medium
surface along-with dried shoot in the center kept in culture
room at 25 ± 2 ºC (1.0 x). 100
5.56: Dried softwood shoot showing no development of roots with
yellowish fungal contamination at base (1.0 x). 100
6.1: Mature bark (arrow) cultured on MS medium supplemented
with 4.52 µM 2, 4-D (2.5 x). 111
6.2: An enlarged view of Fig. 6.1 (3.1 x). 111
6.3: Bark explants cultured on MS medium supplemented with
13.57 µM 2, 4-D showing the induction of watery, translucent
callus (arrows) at day 30 of initial culture (3.1 x). 111
6.4: Mature bark cultured on MS medium supplemented with
22.61 µM 2, 4-D showing the induction of watery, translucent
callus (arrow) at day 26 of initial culture (3.1 x). 112
6.5: Induction of greenish-white, compact callus (arrows) from the
cracked portions (cp) of the mature bark on MS medium
supplemented with 50 µM TDZ, at day 32 of initial
culture (3.1 x). 112
6.6: Creamy-white, watery, translucent, compact callus (arrows)
from the ruptured portions of the bark on MS medium
supplemented with100 µM TDZ, at day 33 of initial culture
(4.0 x). 112
6.7: Yellowish-green, granular, friable and embryogenic callus
on WPM containing 2, 4 D + TDZ (1.0 + 1.0 µM) at day 37
of initial culture (1.3 x). 114
6.8: Formation of compact, transparent and nodular callus from the
cracked portions of the mature bark explants (arrows) cultured
on WPM containing 50 µM TDZ at day 24 of initial culture
(4.0 x). 114
xix
6.9: Mature bark cultured on WPM containing 100 µM TDZ
showing the formation of greenish-yellow, compact nodular
callus (arrows) at day 24 of initial culture (3.1 x). 114
6.10: Greenish-yellow, compact callus (arrows) induced on WPM
containing1.0 µM TDZ at day 33 of initial culture (3.1 x). 115
6.11: Greenish-white, compact callus formed on WPM containing
13.57 µM 2, 4-D at day 30 of initial culture (1.7 x). 115
6.12: Greenish-white, compact callus (arrow) formed on WPM
containing 13.57 µM 2, 4-D at day 57 of initial culture (2.1 x). 115
6.13: Whitish-brown, granular and friable callus on DKW medium
supplemented with 2, 4-D + TDZ (1.0 + 1.0 µM) at day 36 of
initial culture (3.1 x). 116
6.14: Whitish-brown, granular and friable callus on DKW medium
supplemented with 2, 4-D + TDZ (1.0 + 1.0 µM) at day 36 of
initial culture (3.1 x). 116
6.15: Whitish-brown, friable callus on DKW medium supplemented
with 50 µM TDZ at day 39 of initial culture(3.1 x). 116
6.16: Olive-green, compact and nodular callus on DKW medium
containing 4.52 µM 2, 4-D at day 39 of initial culture (2.9 x). 117
6.17-18: Olive-green, compact, nodular callus with white luster on
DKW medium containing 22.61 µM 2, 4-D at day 39 of
initial culture (2.9 x). 117
6.19: Greenish-yellow, compact callus on DKW medium
supplemented with 13.57 µM 2, 4-D (3.1 x). 117
6.20: Greenish-yellow, compact and granular callus indicating
bark remnants (arrow) on DKW medium containing 13.57 µM
2, 4-D at day 39 of initial culture (3.1 x). 117
6.21: Greenish-yellow, compact callus on DKW medium
containing 13.57 µM 2,4-D at day 51 of initial culture (3.1 x). 117
6.22: Greenish-yellow, friable callus on DKW medium containing
100 µM TDZ at day 39 of initial culture (3.1 x). 117
6.23: Whitish-brown, friable callus on DKW medium supplemented
with 1.0 µM TDZ at day 57 of initial culture (3.1 x). 117
xx
6.24: Initiation of browning of greenish-yellow, granular compact
callus on DKW medium supplemented with 100 µM TDZ
after 57 days of initial culture (3.1 x). 118
6.25: Browning of callus after 4th subculture on DKW medium
containing 13.57 µM 2, 4-D (3.1 x). 118
6.26: Browning of callus after 4th subculture on DKW medium
containing 1.0 µM of each 2, 4-D + TDZ (3.1 x). 118
6.27: A) Immature Pecan (Carya illinoensis) fruits attached to a twig,
collected during August 2007 B) An opened view of immature
fruit C) Mature fruits collected during September2007, showing
ruptured outer green husk D) Mature fruits with a pointed tip (T)
and rounded base (B)(arrows) showing outer reddish-brown
hard endocarp with green husk removed E) A longitudinal
view of opened fruit from outside F) A longitudinal view of
opened Pecan fruit cut from the centre into two halves (3.1 x). 119
6.28: Immature Pecan fruit (cotyledonary portion) cultured on
DKW medium supplemented with 50 µM TDZ (3.1 x). 121
6.29: A batch of culture vessels showing cultured fruit parts on
MS medium containing 13.57 µM 2, 4-D (3.1 x). 121
6.30: Yellowish-brown, translucent, watery and compact callus
formed on DKW medium supplemented with 13.57 µM
2, 4-D at day 27 (2.0 x). 122
6.31: Yellowish-brown, smooth and compact callus formed on DKW
medium supplemented with 13.57 µM 2, 4-D at day 39 (3.1 x). 122
6.32: Yellowish-brown, compact and watery callus formed on DKW
medium supplemented with 13.57 µM 2, 4-D after 51 days
(4.0 x). 122
6.33: Yellowish-brown, translucent, watery and compact callus
formed on DKW medium supplemented with 4.52 µM 2, 4-D
(4.0 x). 123
6.34: Yellowish-brown, smooth and compact callus formed on
DKW medium supplemented with 4.52 µM 2, 4-D (2.5 x). 123
6.35: Translucent, granular, watery and compact callus formed
on DKW medium supplemented with 4.52 µM 2, 4-D (4.0 x). 123
xxi
6.36: Off-white, friable, translucent callus observed on DKW
medium containing 22.61 µM 2, 4-D (4.0 x). 123
6.37: Yellowish-brown, compact callus observed on DKW medium
supplemented with 22.61 µM 2, 4-D (3.1 x). 123
6.38: Yellowish-brown, compact, nodular callus observed on
DKW medium supplemented with 31.65 µM 2, 4-D (2.0 x). 124
6.39: Whitish-brown, compact, lustrous callus on DKW medium
containing 31.65 µM 2, 4-D (3.1 x). 124
6.40: Translucent-white, watery callus also showing the greenish
fruit part (arrow) developed on MS medium containing 13.57
µM 2, 4-D (1.7 x). 125
6. 41: Translucent-white, watery callus also showing the greenish
fruit part (arrow) developed on MS medium containing 13.57
µM 2, 4-D (1.7 x). 125
6.42: Transparent white, watery, smooth callus highlighting the
green fruit part (arrow) developed on MS medium containing
4.52 µM 2, 4-D (2.0 x). 125
6. 43: Transparent white, watery, smooth callus highlighting the
green fruit part (arrow) developed on MS medium
containing 4.52 µM 2, 4-D (2.0 x). 125
6.44: Brownish-white, rough, compact callus on MS medium
containing 22.61 µM 2, 4-D (3.1 x). 126
6.45: Brown, smooth, compact callus highlighting the whitish
fruit part (arrow) developed on MS medium containing
22.61 µM 2, 4-D (2.0 x). 126
6.46 - 47: Off-white, rough, compact callus highlighting the greenish-
yellow fruit part (arrow) developed on MS medium
containing 31.65 µM 2, 4-D (2.8 x). 126
6.48: Creamy-white, translucent, watery and smooth callus on
WPM containing 13.57 µM 2, 4-D (3.1 x). 127
6.49: Light yellowish-brown, watery and smooth callus observed
on WPM supplemented with 4.52 µM 2, 4-D (3.1 x). 127
xxii
6.50: Off-white, granular, compact callus on WPM containing
22.61 µM 2, 4-D (3.1 x). 128
6.51: Induction of light-brown, granular, watery callus
(red arrow) highlighting whitish fruit part (black arrow)
on WPM containing 31.65 µM 2, 4-D (3.1 x). 128
6.52: Induction of root primordium (arrow) after callus browning
on DKW medium containing 4.52 µM 2, 4-D (3.1 x). 129
6.53: Induction of root primordium (arrow) after callus browning
on DKW medium containing 13.57 µM 2, 4-D (2.1 x). 129
6.54: Root initiation (arrows) from the callus (after callus
browning had just begun) developed on DKW medium
containing13.57 µM 2, 4-D (3.1 x). 129
6.55: An enlarged view of the Fig. 6.54 highlighting its left
portion. Root initiation (arrow) is quite evident (4.0 x). 129
6.56: Right-side enlarged view from the Fig. 6.54 showing two
roots (arrows) (3.1 x). 129
6.57: A developing root originating from a callus culture on DKW
(4.52 µM 2, 4-D) showing signs of browning at day13 of
induction (4.0 x). 130
6.58: Browning of root primordium (arrows) developed from a
brown callus at day 16 of induction (3.1 x). 130
6.59: Completely necrotic callus at day 110 maintained on DKW
medium containing 13.57 µM 2, 4-D (4.0 x). 130
7.1: Effect of different concentrations of BAP on in vitro shoot
multiplication from immature cotyledon of Pecan on three
different salt formulations, i.e., MS, WPM or DKW. Data
were recorded at day 15 of initial culture. 139
7.2: Multiple shoots (right bracket) developed from immature
cotyledonary portions (arrow) on MS medium
supplemented with 15.0 µM BAP (1.6 x). 140
7.3: Multiple shoots (arrows) originating from immature
embryonic axes (EA with arrow) on MS medium
supplemented with 15.0 µM BAP (1.2 x). 140
xxiii
7.4: Proliferating multiple shoots (right arc) on MS medium
supplemented with 8.0 µM BAP (1.2 x). 140
7.5: Multiple shoots (arrows) proliferating on MS medium
supplemented with 4.0 µM BAP (1.3 x). 140
7.6 - 7.7: Multiple shoot (arrows) induction on MS medium supplemented
with 1.0 µM BAP at day14 of initial culture(2.5 x). 140
7.8: Multiple shoots (left bracket) originating from immature
cotyledonary portions on DKW medium supplemented
with 15.0 µM BAP (1.2 x). 142
7.9: A bunch of multiple shoots (left bracket) originating from
cotyledonary portions on DKW medium supplemented
with 8.0 µM BAP (3.1 x). 142
7.10: Multiple shoots (arrows) proliferating on DKW medium
containing 8.0 µM BAP (1.3 x). 142
7.11: Multiple shoots (arrows) originating from immature
cotyledonary portions on DKW medium supplemented
with 4.0 µM BAP (1.3 x). 142
xxiv
LIST OF TABLES TABLE TITLE PAGE NUMBER NUMBER 4.1: Effect of DKW, MS and WPM medium supplemented
with various levels of BAP on in vitro seed (incised)
germination and other morphological parameters of Pecan
at 25th day of initial culture. 50
4.1a: Analysis of variance for different parameters for in vitro
seed (incised) germination and other morphological
parameters of Pecan at 25th day of initial culture. 51
4.1b: Glasshouse and in vitro germination of Pecan seeds. 51
4.2: Effects of DKW and MS medium with different levels of IAA,
IBA or NAA on rooting in Pecan. 61
5.1: Effects of different environments and media to force epicormic
buds from Pecan logs (40 cm long stem segments) during
winter season (December- January). 78
5.2: Effects of different environments and media to force epicormic
buds from Pecan logs (40 cm long stem segments) during spring
season (February - March). 83
5.3: Effects of different environments and media to force epicormic
buds from Pecan logs (40 cm long stem segments) during autumn
season (August- September). 85
5.4: Analysis of variance for different parameters for shoot forcing
of Pecan logs. 89
6.1: Effect of DKW, MS and WPM medium with different levels
of 2, 4-D, TDZ and NAA on callus induction from mature bark
explants of Pecan. 113
6.2: Effect of DKW, MS and WPM medium with different levels of
2, 4-D on callus induction from immature fruit explants of Pecan. 120
7.1: Effect of different levels of BAP and TDZ supplemented to DKW,
MS or WPM medium on adventitious shoot induction using
immature cotyledonary explants of Pecan. 139
1
CHAPTER 1
INTRODUCTION
The Pecan nut [Carya illinoensis (Wangenheim) C. Koch] is one of the better-
known hickories. It is also called sweet Pecan and belongs to the family “Juglandaceae”.
Pecan is native to North America and also exists in Texas and North of Mexico
(Hancock, 1997; Andersen and Crocker, 2009). In addition, it is grown in Australia,
Brazil, Peru, Israel and South Africa. The U.S is the world’s largest Pecan producing
country. More than 80 % of the world’s Pecan is produced by United States
(Venkatachalam, 2006, 2007). Pecan is very large, deciduous, temperate tree usually
grow 70 - 100 ft in height and 1.0 - 1.5 m in trunk diameter (Ball, 2001), having densities
of 10 - 15 trees per acre (Taylor, 1990). Pecan has adapted to a wide climatic range
between 30 - 42 oN latitude (Sparks, 1991) suggesting noteworthy genetic diversity.
Though Pecan is mostly valued for its commercial nut crop but it has many other uses as
well. An extraction from bark has been used for the treatment of TB (Moerman, 1998). It
also provides food for wild life as Pecan nuts are eaten by a number of birds, fox, gray
squirrels, opossum, raccoons and peccaries (Peterson, 1990). Pecan wood is used for
making agricultural implements, cabinetry, flooring and paneling (Vines, 1960; Little,
2001). Hickories have been known in eastern Asia only since 1912. During the year
1972, Pecan was introduced in Pakistan (Rehman and Jan, 1998). Climatic conditions of
various northern regions of Pakistan are very diverse and favor its growth and
multiplication. Infact, many mature Pecan trees have been identified growing and
fruiting in and around Abbotabad. Although Pecan is an excellent multipurpose tree but
studies for its improvement throughout the world including Pakistan are scarce. There is
to-date a short-fall in Pecan nuts and its products throughout the world because of lack of
2
rapid micropropagation methods and disease attacks. Considering the significance of
Pecan industry it is desired that micropropagated Pecans may be grown on mass-scale in
Pakistan also.
Generally, Pecan is propagated through seeds. Seed germination may be defined
as “the resumption of active growth in an embryo which results in its emergence from
the seed and development of those structures essential to plant development” (Bonner,
1984). Pecan seeds show delayed germination as outer hard shell physically hampers the
emergence of the radicle. In vitro seed germination holds promise to enhance the
germination potential in Pecan. Moreover, in vitro studies can provide insights into in
situ plant responses to external environment and basic information of early plant growth
and development (Dutra et al., 2008). To the best of our knowledge, there are no prior
reports on extensive in vitro seed germination trials of Pecan. However, in vitro seed
germination has been reported in several other plant species (Maliro and Kwapata, 2000).
The use of various growth regulators has also been reported for in vitro seed germination
(Nikolic et al., 2006). This practice hold great promise to overcome sexual complexities
associated with propagation of plants using conventional means. Another method for the
vegetative propagation of most of the plants is clonal propagation. Budding and grafting
are the most popular methods for Pecan’s propagation and its many horticultural
varieties are propagated by these methods (Smith et al., 1974; Peterson, 1990).
Vegetative propagation via rooting of mature cuttings could produce only a few
propagules that were inadequate for rapid clonal multiplication of Pecan. Clonal Pecan
rootstocks however have been achieved with only limited success (Wolstenholme and
Allan, 1975).
Tissue culture techniques offer unique opportunities for the rapid multiplication of
many plants. In vitro propagation studies of woody plants have shown that these
3
techniques offer a solution to problems associated with their rapid propagation (Bonga
and Durzan, 1987; Ahuja, 1991). However, woody plants are generally recalcitrant to in
vitro regeneration (Benson, 2000). Attempts at Pecan tissue culture were reported by
several workers (Smith, 1977; Knox, 1980) but neither was successful in obtaining plants
established in soil. However, Hansen and Lazarte (1982) were successful in the rooting
of juvenile Pecan shoots obtained in vitro. Micropropagation has become a fundamental
aspect of scientific studies and commercial propagation in many plants (Zimmerman et
al., 1986; Dirr and Heuser, 1987) and its advantages as propagation has been described
by several others (Debergh, 1987; Pierik, 1999; Hartman et al., 2002; Debnath et al.,
2006). Softwood shoot forcing is relatively a newer approach for micropropagation and
its potential has been reviewed extensively by Preece and Read, (2003). Shoot forcing
was primarily focused on the use of shoot tips harvested from trees and shrubs during the
dormant season (Read and Yang, 1991). Large branches excised from juvenile portions
of the intact trees and shrubs can also be used to force softwood shoots (Cameron and
Sani, 1994; Henry and Preece, 1997a, b). The forced softwood shoots can be rooted
using general nursery practices (Henry and Preece, 1997a) or as explant source for in
vitro studies (Van Sambeek et al., 1997a; Preece and Read, 2003). Apart from several
other temperate woody species, shoot forcing as well as forcing large stem segments has
never been attempted before in Pecan. However, Aftab and Preece (2007) discussed the
possible extension of these methods in Pecan. Besides micropropagation, an alternative
technique with potential application to mass propagation is somatic embryogenesis
(Rodriguez and Wetzstein, 1994). This aspect though needs further research.
Adventitious regeneration or shoot organogenesis is another technique that holds
promise for whole plant regeneration. The information on adventitious shoot
regeneration in Pecan is quite limited in the contemporary literature though callus
4
formation and plantlet regeneration has been reported in Pecan (Corte-Olivares, et al.,
1990b; Yates and Reilly, 1990). Organogenesis in Pecan has been reported from
immature embryonic axes (Yates and Wood, 1989) and mature cotyledons and
embryonic axes (Obeidy and Smith, 1993). Shoot organogenesis and plantlet formation
has been well documented in various other trees (Pandey and Jaiswal, 2002; Lyyra, et
al., 2006; Rajeswari and Paliwal, 2008). Thidiazuron (N-phenyl-N'-1, 2, 3-thidiazol-5-yl-
urea; TDZ), a non-purine cytokinin-like compound, has shown promise for in vitro
studies in recalcitrant woody plants. It exhibits stronger effects than conventional
cytokinins over a wide range of species (Curcuma longa, Calendula officinalis,
Azadirachta indica, etc). Its potential use in micropropagation seems to be extending
from the woody plant species to other plant groups as well. Therefore, TDZ was
employed to demonstrate its role in tissue culture studies of Pecan.
The present study was undertaken with the objective to establish successful
protocols for micropropagation, callus induction, plant regeneration and acclimatization.
Considering its scarce resource in Pakistan, the current investigation also aims to
produce an adequate amount of Pecan stock. For this purpose certain novel or newer
micropropagation methods such as shoot forcing or forcing large stem segments were
also investigated for Pecan’s multiplication. It would hence extend our knowledge about
various aspects of in vitro growth and differentiation in Pecan.
5
CHAPTER 2
LITERATURE REVIEW
In the words of Read and Paek (2007), “modern biotechnology owes much to its
roots derived from plant tissue culture and micropropagation”. Gottileb Haberlandt
(1902) is referred to as the “Father of Tissue Culture”, is often cited as the “origin and
emergence of plant tissue culture and its subsequent application”. Plant tissue culture
techniques have become a fundamental tool for studying and solving basic and applied
problems pertaining to agriculture, industry, environment and health in plant
biotechnology. These techniques have greater impetus in the field of propagation (Islam,
1996). Plant tissue culture is multi-dimensional field that offers excellent prospects for
plant improvement and crop productivity (Jain, 2001). Since the establishment of
cultivation of plants, mankind is looking for methods that aid in the mass multiplication
of plants using minimum quantity of propagules. The ultimate result of their enquiry
leads to the development of tissue culture techniques. Woody plants having economic
significance are generally propagated by seeds. Propagation of plants through tissue
culture has become an essential and popular technique to reproduce crops that are
otherwise difficult to propagate conventionally by seed and/or vegetative means.
Grafting and budding are the other conventional methods of vegetative propagation. Due
to several limitations in conventional propagation methods certain relatively newer tissue
culture techniques were developed for tree improvements. Different plant parts such as
apical meristem, nodal explants, cotyledons or leaf explants were used for
micropropagation of woody trees. For multiple shoot induction, cotyledonary nodal
explants have been used in tree propagation (Das et al., 1996; Pradhan et al., 1998; Das
et al., 1999; Purohit et al., 2002; Walia et al., 2003). Genetic variations during callus
6
cultures and micropropagation of trees have also been reported (Gupta and Varshney,
1999). Some molecular markers such as RAPD have also been used to detect genetic
variations among in vitro clones (Gangopadhyay et al., 2003).
2.1 TISSUE CULTURE STUDIES IN PECAN
Pecan is a native hardwood tree species in US, mostly grown for its edible nut
with high commercial value. The first known selections were made in 1846, and many
cultivars were available by the late 19th century (Madden and Malstrom, 1975).
Recently, more than 500 Pecan cultivars, each with unique traits were documented in
literature (Andersen and Crocker, 2009) but only four of them have become the
standards of the Pecan industry. These include Stuart, Desirable, Western Schley and
Wichita. Pecan cultivars are usually propagated through seeds. It has been shown by
several workers that the larger nuts of Pecan make larger seedlings (Adams and Thielges
1977; Herrera and Martinez, 1983) hence sizing of nuts may be beneficial. Budding and
grafting have been the primary means of improvement, but newer studies (Grauke et al.,
1990) and research on the reproductive biology and genetics of Pecan (Graves et al.,
1989; McCarthy and Quinn, 1990; Yates and Reilly, 1990; Yates and Sparks, 1990)
demonstrates the promise for future improvements in nut production and disease
resistance. In vitro studies for Pecan improvement throughout the world are generally
scanty. Tissue culture techniques have been developed for several tree crops, but
previous efforts with Pecan have shown that it is difficult to propagate by in vitro
methods (Wood, 1982). These techniques have been used in Pecan mainly for the
purpose of clonal propagation.
Previously, various aspects of research on Pecan includes; studies on propagation
(Smith et al., 1974), seed germination and dormancy (Dimalla and Van Staden, 1977),
7
micropropagation (Hansen and Lazarte, 1984), seed maturation and germination (Wood,
1984), adventitious regeneration (Long et al., 1995), cell suspension cultures (Burns and
Wetzstein, 1997), somatic embryogenesis (Rodriguez and Wetzstein, 1998), manganese
deficiency (Smith et al., 2001), effect of Zinc supply on growth and nutrient uptake (Kim
et al., 2002a), effect of nitrogen form and nutrient uptake (Kim et al., 2002b), forcing
shoot tips and epicormic/ latent buds (Preece and Read, 2003) and use of Pecan nutshells
as biosorbent for the removal of toxic metals from aqueous solutions (Vaghetti et al.,
2009). Recently, embryos from immature fruits of Pecan were germinated in vitro
(Payghamzadeh and Kazemitabar, 2010).
In this section a brief review of work is given in a manner so as to highlight the
contemporary status of the research work in Pecan tissue culture.
2.1.1 MICROPROPAGATION
Micropropagation is the complex blend of science and art (Lineberger, 1981;
McCown and McCown, 1999). As a concept, micropropagation was first presented to the
scientific community in 1960 by Morel producing virus-free Cymbidiums.
Micropropagation is a sophisticated technique for the rapid and large-scale propagation
of many tree species (Chand and Singh, 2004). It has a great commercial potential due to
extremely high speed of multiplication, the high plant quality and the ability to produce
disease-free plants. Micropropagation has been applied to several woody tree species
(Bonga and Aderkas, 1992). Generally, woody plants are recalcitrant to in vitro
regeneration (McCown, 2000; Munshi et al., 2004). The pertinency of micropropagation
for woody trees has been confirmed feasible since the aspects of the system have
established that trees produced by this method are similar to those produced by
traditional methods (Lineberger, 1981). Furthermore, Lineberger (1981) however,
8
described that “the major impact of plant tissue culture will not be felt in the area of
micropropagation, however in the area of controlled manipulations of plants at the
cellular level”.
Many workers have reported propagation of Pecan through conventional methods
(Smith et al., 1974; Brutsch et al., 1977). However these methods suffer various
limitations thus provide few propagules from selected individuals. Several efforts at
Pecan tissue culture were reported by Smith (1977) and Knox (1980) but neither was
successful in establishing plants in soil. However, Knox (1980) obtained few shoots and
plantlets when inverted nodal cuttings were used in vitro which upon transplanting did
not survive. Later, Knox and Smith (1981) successfully proliferated in vitro axillary
shoots of Pecan using seedling explants. Success was limited to the formation of callus
with only few shoots and root formation. Major drawbacks to clonally propagate Pecan
are the poor rooting and their survival rate after transplanting to greenhouse (Brutsch et
al., 1977).
In 1982, Wood successfully induced shoot proliferation in axillary buds of nodal
explants and reported that synthetic hormones with combination of 4.0 mg/ litre BA and
1.0 mg/ litre IBA were most effective for shoot proliferation. Gibberellic Acid (GA3) at
3.0 mg/ litre plus 0.1 mg/ litre BA also enhanced shoot elongation although he was
unable to subculture shoots and rooting was not achieved. In another work performed by
Hansen and Lazarte (1982) shoots were proliferated from juvenile Pecan in vitro and
limited success was reported in terms of rooting.
Hansen and Lazarte (1984) obtained single node cuttings from 2-month-old Pecan
seedlings and induced bud break to from multiple shoots on liquid WPM and 2 %
glucose supplemented with 3.0 mg/ l 6-Benzylamino purine (BA). The shoots developed
in vitro adventitious roots and showed vigorous root system with profused lateral
9
branching from primary roots on transferring to soil after soaking in 10 mg/ l IBA for 8
days. Etiolation of stock plants did not improve shoot proliferation or rooting under in
vitro culture.
Corte-Olivares and co-workers (1990a) reported a procedure for propagating
Pecan using explants from adult trees. They collected nodal explant material during
two consecutive seasons from grafted ‘Western Schley’ trees. Specific trees
representing the vegetative phase, partially bearing phase and fully bearing phase were
identified and three collections of axillary buds were made from them each year. Buds
were cultured on Dunstan and Short (1977) basal medium supplemented with 0.51 mM
ascorbic acid and 4.4 µM BA. They found severe contamination problems which
resulted in the data that were not amenable to statistical analysis in five of six
collections of explants. Even so, in one of these six collections, shoot development and
multiplication was observed during second and third culture passages from transitional
tree explants and from juvenile tree explants in the fourth collections. Amenable data
were found in one of six collections where explants of all three-donor tree phase
responded with shoot multiplication. The results of this preliminary study indicated
that selected adult phenotypes had a potential for clonally micropropagating Pecan.
2.1.2 SOMATIC EMBRYOGENESIS
Somatic embryogenesis has been known in tissue cultures of a wide range of
higher plants, including both angiosperms and gymnosperms (Halperin, 1995). Somatic
embryogenesis is a valuable tool of interest in plant biotechnology for its potential
applications in clonal propagation, genetic transformation and studies involving embryo
development. In addition, somatic embryogenesis is also used for regenerating transgenic
trees. It involves the development of somatic cells into embryos, which proceeds through
10
a sequence of morphological stages that resemble zygotic embryogenesis (Dodeman et
al., 1997; Dong and Dunstan, 1999). In vitro induction of somatic embryogenesis under
controlled environment offers a possibility to study the developmental pathways leading
to somatic embryogenesis (Visser et al., 1992).
Somatic embryogenesis has been reported in several temperate and tropical tree
species (Jain and Gupta, 2009). It is reported that many species of tropical fruit trees
could produce somatic embryos in tissue culture (Litz, 1985). In other studies, temperate
fruit species including apple, sweet cherry, grapes, guava etc. have also been reported to
produce somatic embryos (Tisserat et al., 1979; Ammirato, 1983; Rai et al., 2007). A
successful somatic embryogenesis has been reported in members of the Pecan (Carya
illinoensis) family (Juglandaceae), i.e., Juglans nigra, Juglans hindsii using immature
zygotic embryo explants (Tulecke and McGranahan, 1985). However, the application of
somatic embryogenesis for the improvement of Pecan is still limited as a result of
problems with low initiation frequencies, maintenance of embryogenic cell lines and low
conversion rates.
Somatic embryogenesis is best known as an alternative pathway to propagate
Pecan via methods of tissue culture mainly due to high multiplication rates, formation of
organized root and shoot axes and feasibility of mechanization. A number of studies
have focused on Pecan somatic embryogenesis and conversion to complete plantlets
(Merkle et al., 1987; Wetzstein et al., 1988; 1989; 1990; Corte-Olivares et al., 1990b and
Yates and Reilly, 1990). Somatic embryogenesis has been used for induced regeneration
from in vitro tissue culture, occurring indirectly from callus, cell suspension, or
protoplast culture or directly from cells of an organized structure such as stem segment
or zygotic embryo (Williams and Maheswaran, 1986). They also described the
fundamental homologies between direct and indirect somatic embryogenesis and
11
between single cell and multiple cell initiation. The observed pattern of morphogenesis
depends whether a group of cells establish and maintain coordinated behavior and
influenced by factors, which affect intercellular communication. McGranahan et al.,
(1987) obtained genetic transformation using somatic embryogenic cultures in Juglans.
Wetzstein et al., (1996) suggested that somatic embryogenesis has the potential for
propagating Pecan rootstocks and useful in introducing genes of commercial interest.
Merkle et al., (1987) induced somatic embryogenesis from immature zygotic
embryos of Pecan cultivars “Stuart” and “Desirable”, within one month following
transfer from modified WPM with 2.0mg/ litre 2, 4-D and 0.25 mg/ litre BA in the light
to hormone-free medium in the dark but with low embryogenic frequency. Wetzstein and
co-workers (1988) however, improved the embryogenic frequency up to 40 % for some
explants sampling stages of Pecan.
In another study, Wetzstein and co-workers (1989) examined the effect of
cultivars, sampling date, tree source of explants and duration on conditioning medium
for the optimum production of somatic embryos in two cvs. (‘Stuart’ and ‘Desirable’) of
Pecan. Significant variations in embryogenic response were observed in both the
cultivars. A short term exposure to 2, 4-D was shown to be quite adequate for
embryogenesis in Pecan. Immature zygotic embryos collected in a developmental stage
of rapid cotyledon expansion showed highest embryogenic response, i.e., 54.7 % in
Desirable and 85.2 % in Stuart. No significant effect of duration on conditioning medium
on embryogenic response was observed in both the cultivars. In Stuart, effect of different
trees as explant sources was not significant but found significant in Desirable. However,
plant regeneration and transplantation remained a limiting factor.
Later, Corte-Olivares and co-workers (1990b) reported the induction of somatic
embryogenesis in two cultivars (‘Western Schley’ and ‘Wichita’) with low
12
developmental frequencies into complete plantlets. Growth regulators with different
combinations had a significant effect on induction of embryogenic callus. They proved
that medium containing 2, 4-D was most effective for the induction of embryogenesis.
The individual shoots isolated from shoot multiplication cultures were rooted with 49 %
frequency upon culture for 4 weeks on BDS (Dunstan and Short, 1977) basal medium
containing 14.8 µM IBA. Their results indicated the potential to successfully obtain
complete plants from Pecan somatic embryos.
Studies of Yates and Reilly (1990) on relation of cultivar’s response on somatic
embryogenesis and subsequent plant development revealed that explants of micropylar
region when removed from fruits in the liquid endosperm stage were more embryogenic
than the intact ovules. Medium containing auxin alone or auxin and cytokinins produced
more somatic embryos than medium containing cytokinin alone.
Furthermore, Wetzstein et al., (1990) examined effects of zygotic embryo
explanting time and auxin type on somatic embryogenesis during conditioning in Pecan
(Carya illinoensis). Maximum embryogenesis was observed after 15 weeks post
pollination. Percent somatic embryogenesis and embryo form was significantly affected
by auxin type and concentration but not the embryogenic efficiency. MS medium proved
to be better than WPM for embryo germination.
In another interesting study, Mathews and Wetzstein (1993) established new
methods to increase plant regeneration by repetitive secondary embryo formation which
can efficiently produce large number of clonal plants suitable for establishment in
greenhouse. Silver nitrate (29.43 µM) incorporation to WPM and application of 6-
benzylaminopurine (100 µM) on shoot apices increased maximum shoot regeneration
frequency with average frequency (20 %) of plantlet conversion up to a maximum of 71
13
% in cv. Mahan. Later, 70 - 80 % of the regenerated plants attained hardening stage and
> 99 % of hardened plants were established successfully in the greenhouse.
Later, Rodriguez and Wetzstein (1994) investigated callus production, embryo
formation and embryo morphology in Pecan. Explants were cultured for one week on
WPM with either NAA or 2, 4-D at a concentration of 2, 6 or 12 mg/litre and then
subcultured on fresh basal medium. The best auxin treatment was 6 mg/litre NAA in the
induction medium, with 100 % somatic embryogenesis in cv. Stuart. Somatic embryos
induced by NAA were shown to have relatively normal morphology than those induced
by 2, 4-D. They reported that somatic embryo morphology affects plantlet conversion
and NAA proved to be a superior auxin than 2, 4-D for the production of somatic
embryos and their subsequent conversion to plants.
In 1998, Rodriguez and Wetzstein critically compared morphological and
histological aspects of Pecan somatic embryos induced on media with NAA or 2, 4-D.
The media containing NAA or 2, 4-D had shown significant differences in the timing and
pattern of initiation and development of somatic embryos. Embryos derived from callus
cultures on NAA had normal morphology while those derived from cultures on 2, 4-D
had higher incidences of abnormalities. Their study strongly revealed the multicelluar
origin of embryos in contrast to earlier studies of somatic embryogenesis where embryos
were defined as having single-cell origin (Street and Withers, 1974).
Yates and Wood (1989) demonstrated organogenesis from immature embryonic
axes in vitro in Pecan. Highest number of normal plants was produced from medium
containing IBA, BA and kinetin at 0.5, 4.4 and 9.3 µM respectively. Shoots only were
produced on a medium containing cytokinins and rooting was observed on medium with
no cytokinins. In cv. ‘Desirable’ greatest number of axillary shoots were elongated from
14
embryo axes on a medium containing cytokinin only, but both with auxin and cytokinins
for cv. ‘Stuart’.
Later, Obeidy and Smith (1993), investigated organogenesis from mature Pecan
cotyledons and embryonic axes. Embryonic axes at cotyledonary nodes formed 85 %
microshoots and 30 % were rooted on an auxin-free medium after pre-culture in a
medium with 20 µM IBA. Adventitious buds emerged on callus surface previously
produced on medium containing TDZ (25 µM) from cotyledonary nodes and radicals.
Kumar and Sharma (2005) induced somatic embryos from cotyledon explants of
Walnut and Pecan. They cryopreserved these somatic embryos using non-toxic
cryoprotectants, i.e., DMSO, glycerol and ethylene glycol and evaluated their survival
percentage. Maximum survival percentage was observed with 5 % DMSO, 1.5 %
glycerol and 3 % ethylene glycol pre-treatment. In contrast, higher sucrose levels
decreased survival rate and the embryos became necrotic. However, sucrose-desiccated
somatic embryos pre-treated with cryoprotectants survived better after one day in the
liquid nitrogen.
Somatic embryogenesis can be applied for efficient plant regeneration systems. It
may also be utilized for introducing the genes of interest. Molecular markers can be used
as a means of evaluating genetic stability of plants regenerated through tissue culture.
Somatic embryos exhibit morphological features similar to zygotic embryos. Abnormal
developments, however, have frequently been observed and genetic fidelity of embryos
hence becomes questionable. Therefore, the genetic fidelity of culture must be evaluated
before somatic embryogenesis can be exploited. In one such interesting research work,
Vendrame et al., (1999) evaluated the applicability of using AFLP analysis to assess the
genetic variability in somatic embryos of Pecan (Carya illinoensis) and compared
between and within embryogenic culture lines. They revealed that individual embryos
15
derived from the same culture line exhibited high similarity and could be grouped
together. However, within a culture line some embryo-to-embryo differences were also
observed. They concluded that AFLP can be used as a reproducible technique to check
the genetic variation among Pecan somatic embryo cultures. Larkin and Scowcroft
(1981) were the first who designated variations in tissue-culture-derived plants as
somaclonal variations. Somaclonal variations were also detected in Peach regenerants
when developed from two different embryo callus cultures using RAPD (Hashmi et al.,
1997). They suggested that genetic changes occurred during tissue culture. Brown et al.,
(1993) were also successful in genetically distinguishing among wheat suspension
culture lines and also among regenerated plants through RAPD.
Several studies have been reported to the use of molecular markers in
understanding the Pecan genome. The genetic diversity of Pecan populations through
isozyme system has been demonstrated by Marquard 1987, 1991; Marquard, et al., 1995;
Ruter et al., 2000, 2001. Conner and Wood (2001) employed RAPDs for the
identification of Pecan cultivars and estimated their genetic relatedness. The molecular
evaluation of Pecan trees regenerated from somatic embryogenic cultures was carried out
by Vendrame et al., (2000) using AFLPs. Grauke et al., (2001) reported mean 2C
genomic size of Pecan to be approximately 1.7 pg. Later, in another study, Grauke et al.,
(2003) evaluated simple sequence repeat (SSR) markers for the genetic study of Pecan.
Crespel et al., (2002) stated that molecular markers are valuable in perennial crops for
the construction of linkage maps. Molecular linkage maps are successfully employed in
many crops for directed germplasm improvement (Pearl et al., 2004). Recently,
molecular linkage maps of several tree fruit and nut crops have also been produced,
including Pear (Yamamoto et al., 2002), Apricot (Lambert et al., 2004) and Walnut
(Fjellstrom and Parfitt, 1994). In one such work, Beedanagari et al., (2005) reported a
16
first genetic linkage map of Pecan using RAPD and AFLP markers. These maps were
considered important towards the detection of genes controlling horticulturally important
characters such as nut size, maturity date, kernel quality and disease resistant (Conner,
1999).
To initiate further work on Pecan, somatic embryogenesis has also been
attempted by using cell suspension cultures. Regenerable suspension cultures established
an attractive tool for the production of clonal plants and in studies involving genetic
transformation. Previously, repetitive somatic embryogenesis was first reported in Pecan
(Merkle et al., 1987) on solidified medium. Later, a number of research workers have
improved the quantity (Wetzstein et al., 1989; Yates and Reilly, 1990) and quality
(Wetzstein et al., 1990) of the somatic embryos through modified culture media and
conditions. Even through many modifications of the cultured media, none of any prior
information illustrated any system for the production and development of somatic
embryos in liquid culture medium. In liquid suspensions, synchronized development of
the embryogenic cultures was one of the major advantages over the solidified cultures.
In tissue cultures of Pecan, stable embryogenic suspensions have been developed
by Burns and Wetzstein (1994). They induced pre-globular stage embryo masses on
hormone-free liquid suspension cultures of Pecan to develop into somatic embryos on
semi-solid medium. Effect of modified solid medium (various combinations of ABA,
Maltose, casein hydrolysate and filter paper overlays) treatments on somatic embryo
storage reserve accumulation was investigated. Embryos analyzed for triglycerides and
protein contents showed significant reserve deposition for some treatments but
associated with undesirable deterioration in embryo morphology. The treatment that
enhanced the reserve accumulation was identified promoting plant recovery from
suspension-derived Pecan somatic embryos.
17
Later, in another interesting work, Burns and Wetzstein (1997) developed a
method for the establishment and proliferation of developmentally stable, embryogenic
Pecan suspension cultures, presenting a major improvement in embryogenic tissue
culture in Juglandaceae. The established suspension cultures consisted of a mixture of
pre-globular, globular stage embryo aggregates and freely suspended globular embryos.
Their studies revealed that cultures were repetitively embryogenic and proliferated in
growth-regulator-free medium. Repetitive somatic embryogenic cultures have also been
reported in some other related member of the family Juglandaceae such as; Juglans regia
(Tulecke and McGranahan, 1985) and Juglans nigra (Neuman et al., 1993; Preece et al.,
1995).
2.1.3 NOVEL MICROPROPAGATION METHODS
Previous tissue culture work involved micropropagation of cuttings obtained
from seedlings or buds of trees grown under field conditions. The rooting of these shoots
was slow or altogether not possible. On the other hand, contamination was another major
constraint encountered when these shoots are used for in vitro cultures. Shoots taken
from outdoor usually have microbes in tiny cracks of bark, not removed through
disinfestation causing in vitro contamination of cultures (Preece and Read, 2003).
Therefore, some other relatively newer techniques have been developed that utilize the
parts of the plants (branch tips and/ or stem segments) during dormant season and force
new growths in a greenhouse environment. These techniques, such as shoot forcing as
well as forcing epicormic buds may provide a breakthrough in the micropropagation of
woody plants as well as for herbaceous species. These forcing techniques also have the
potential for commercial propagation of plants. Research has been conducted on shoot
forcing for years but much focus was on shoot tips harvested from trees and shrubs
during the dormant season (Read and Yang, 1991). For softwood shoot forcing, shoot
18
tips of specific length (20 - 25 cm long) were cut, surface disinfested and placed in a
solution containing 8- hydroxyquinoline citrate (8-HQC) and different growth regulators
(Yang and Read, 1992, 1993). On the other hand, large branches (40 cm long) excised
from juvenile portions of the trees and shrubs can also be used to force softwood shoots
on a greenhouse media (Cameron and Sani, 1994, Henry and Preece, 1997a, b). No
forcing solution is used in this technique. These forced softwood shoots can be rooted as
stem cuttings (Henry and Preece, 1997a). Softwood shoots can also be utilized as an
explant source for in vitro studies and micropropagation (Preece, 2003).
Clonal propagation is achieved by culturing nodal explants taken from in vitro
seedlings or form field-grown adult trees. Hence, for in vitro establishment of softwood
shoots, there is a need to obtain explants with minimum of contamination. Read and
Yang, (1988) disinfested the shoot tips treating with a solution of 0.78 % NaOCl
containing Tween-20. Shoot tips were forced by placing in a forcing solution containing
BA and GA3. They reported that the use of GA3 favored bud break and consequently
increased multiple shoot production under in vitro conditions.
Read and Yang (1991) later forced softwood shoots from privet (Ligustrum
vulgaris) and arrowwood (Viburnum dentatum) and tested different growth regulators in
forcing solution for rooting of softwood cuttings. They reported that IBA increased
number of roots per cuttings for both plants while root length increased only in Privet.
On the other hand, GA3 decreased number of roots per cutting as well as reduced root
length.
Similarly, in another study, Yang and Read (1992) reported the influence of pre-
forcing treatment on bud break and shoot elongation of lilac, Privet and Vanhoutte
spirea. Their results revealed that pre-forcing treatments increased bud break by 20 %
19
and shoots were elongated 3.0 mm greater as compared to control. However, pre-
treatment effect differed with the plant species.
In 1993, Yang and Read forced Vanhoutte spirea stems in forcing solution
containing 8- hydroxyquinoline citrate (8-HQC), 2 % sucrose with different levels of BA
and GA3 to observe their effects on in vitro cultures. They revealed that LS (Linsmaier
and Skoog, 1965) medium supplemented with 5 µM BAP or 5 µM BAP + 1 or 5 µM
IAA was superior for the shoot forcing in “Vanhoutte spirea”. Addition of BAP to
forcing solution enhanced shoot proliferation while GA3 reduced shoot establishment in
vitro.
Large stem segments having epicormic (dormant, latent or suppressed) buds cut
during the dormant season can also be forced by placing in a suitable glasshouse
medium. Large numbers of epicormic buds are present on stems of several woody tree
species. Softwood shoots developed from epicormic buds on large stem segments can be
used as stem cuttings in nursery industry (Cameron and Sani, 1994; Henry and Preece,
1997b).
Henry and Preece, (1997a) investigated the production of softwood shoots and
their subsequent rooting in maple species. The percent softwood shoot production varied
considerably within the species and clones of genus Acer. However, greater (59 %)
number of softwood shoots was rooted in red maple as compared to either in sugar (15
%) or Japanese maple (26 %). Furthermore, Henry and Preece, (1997b) studied the
influence of length and diameter of large stem segments on the production of softwood
shoots from epicormic buds of selected species of genus Acer. They concluded that both
stem length and diameter influenced the production of softwood shoots. Their study
revealed that stem segments ranging from 30 - 40 cm long with 5.2 - 7.6 diameters were
best for the softwood shoot production.
20
Preece et al., (2002) developed a system for the production of softwood cuttings
during the dormant season. It provided a longer growing season to force and root
softwood segments in mid to late winter during the year of propagation for plant growth,
hence proved advantageous over traditional propagation methods. They suggested
intermittent mist to be the most effective forcing environment. Juvenility seems to be an
important factor and it is easier to propagate plants in the juvenile growth stage than the
adult phase. Similarly, microshoots originated from adult black walnut were hard to root
than that of juvenile origin (Heile-Sudholt et al., 1986).
Van Sambeek et al., (2002) forced branch segments of adult hardwoods for
production of softwood cuttings from the latent buds under greenhouse conditions. A
maximum of 10 - 15 visible buds were sprouted and elongated to produce softwood
shoots during February to April. They also reported sugar maple to be least productive
failing to induce fewer sprouts per meter of branch wood than that of other twelve
hardwood species assessed. In addition, intermittent mist throughout the day was more
successful than continuous mist for forcing epicormic buds.
In 2003, Preece and Read reviewed two novel methods for micropropagation i.e.,
forcing softwood shoots using forcing solution and/ or forcing large stem sections in
greenhouse media in many woody plants. Neither technique was used widely at that time
but appeared to have great potential for woody plant micropropagation. They reported
the possibility of shoot forcing by cutting 20 - 25 cm of shoot tips and placing in a
solution containing 8- hydroxyquinoline citrate (8-HQC), 2 % sucrose and different
growth regulators. In order to force softwood or epicormic shoots, branches from
juvenile tree portions were cut into sections (30 - 35 cm long) and placed horizontally in
flats or benches filled with perlite. The forced soft wood shoots were excised, surface
disinfested and used as explants for micropropagation. The use of intermittent mist was
21
the best forcing environment. However, they noted that chances of microbial
contamination usually existed for softwood shoots forced under intermittent mist and
used in vitro. They suggested that to minimize microbial contamination, watering should
be in such a way to have no direct contact with the sprouts. Careful manipulation
resulted in successful establishment of softwood shoots-derived explants in vitro (Van
Sambeek et al., 1997a, b).
In another research work involving silver maple (Acer saccharinum) and green
ash (Fraxinus pennsylvanica), Aftab et al., (2005) reported the effect of three
environments (lab, mist or fog) four media (perlite, vermiculite, 1 perlite: 1 vermiculite
by volume) and H2O2 treatments on shoot forcing and subsequent transfer of explants
derived from forced epicormic buds under in vitro conditions. A significant interaction
was observed among perlite, vermiculite and environment with the most shoots (6.7/
stem segment) produced under mist. Explants from in vitro cultures had only 4 %
microbial contamination as compared to explants from mist (92.2 %). They found that
with the application of Zerotol, contamination decreased to 43 % and 46 % clean
explants were established when stems were placed under mist and drenched weekly with
0.18 % H2O2.
Later, in another study, Preece and Read (2007) forced leafy explants and
cuttings from the woody species. They demonstrated that stem diameter and stem length
significantly influenced the softwood shoot production in woody species.
Previously, mostly temperate woody species have been forced from large stem
sections (Preece et al., 2001; Preece et al., 2002; Van Sambeek et al., 2002). Aftab and
Preece (2007) studied forcing and in vitro establishment of temperate (silver maple,
green ash or Pecan) as well as for the first time in tropical tree i.e., Tectona grandis
(Teak). They got 6.7 shoots per stem segments in silver maple when forced under mist
22
on perlite/ vermiculite medium. Green ash produced 1.2 mean number of softwood
shoots. In Pecan, microbial contamination was the major limiting factor for softwood
shoot production and establishment in vitro. However, they obtained 3 mean numbers of
shoots per log when forced under lab conditions. On the other hand, 5 mean numbers of
shoots were produced in teak under glasshouse conditions on sterilized sand.
Later, in another study involving teak (Tectona grandis L.), Akram and Aftab
(2009) reported an efficient method for clonal propagation and in vitro establishment of
softwood shoots from epicormic buds developed under light or shade conditions. Higher
numbers of softwood shoots (6.17) were produced under light conditions as compared to
shade. The length of softwood shoots, number of nodes and leaves were also higher
under light conditions. Their results revealed autoclaved sand to be best forcing medium
for teak. Shoot apices (60 %) and nodal explants from softwood cuttings were
successfully established in vitro and afterwards acclimatized to greenhouse conditions.
Many researchers have been working constantly to improve this technique, i.e.,
forcing softwood shoots from large stem segments. Recently, Mansouri and Preece
(2009) investigated the effect of various levels of growth regulators on softwood shoot
production from large stem segments of Acer saccharinum. They used BA and/ or GA3
in the latex paint and painted the large stem segments. Softwood shoots initiated on the
stems painted with 3 mM BA, produced greater number of shoots (4.3) when cultured on
medium supplemented with 0.01 µM TDZ as compared to control or other
concentrations of BAP used. Callus formation was also observed frequently from the
stem explants treated with 3 mM BA and transferred to medium containing 0.01 µM
TDZ. They suggested that stem segments treated with PGR’s in latex paint expands the
season to grow softwood shoots throughout a longer period of the year that can be
utilized as explants source in vitro.
23
Moreover, most of the studies on the use of epicormic buds as a source of
juvenile explants have been conducted with trees, it can also be applied to shrubs as
established by Pereira (2009). He developed an in vitro method for the propagation of
shoots by axillary bud proliferation on epicormic stems in Vaccinium cylindraceum Sm.
He demonstrated softwood shoot forcing as a reversion to juvenility in mature wild
Shrub Vaccinium cylindraceum. They also demonstrated that shoots derived from
epicormic buds have juvenile morphological characteristics and can be utilized for
micropropagation studies.
Even though, most of the previous work has been conducted on several temperate
woody tree species such as silver maple, red maple, Japanese maple, green ash and
privet, softwood shoot forcing was not attempted before in Pecan (Carya illinoensis).
However, it was anticipated that possibilities to force softwood shoots from epicormic/
latent buds also exists with the Pecan (Aftab and Preece, 2007). During their work, high
microbial contamination was the major limitation for establishment of Pecan under in
vitro condition. However, they suggested that this technology is quite promising for the
propagation of this recalcitrant tree species.
2.1.4 ADVENTITIOUS REGENERATION
Adventitious regeneration means the production of adventitious shoots and buds
from tissues other than axillary buds, e.g., leaves, hypocotyls and the cotyledonary
explants. The most common explant used for adventitious regeneration of woody plants
is cotyledons (Huetteman and Preece, 1993). They may either be from mature or
immature seeds and leaf tissue from in vitro cultures. However, adventitious regeneration
has also been achieved by using various explants such as leaf tissues of Prunus dulcis
(Ainsley et al., 2000), young leaves from in vitro-grown shoots of Phellodendron
24
amurense (Azad et al., 2005), hypocotyls of Feronia limonia (Vyas et al., 2005),
immature cotyledons of Prunus persica (Wu et al., 2005), Prunus dulcis (Ainsley et al.,
2001) and mature stored cotyledons of Prunus sp. (Canli and Tian, 2008; 2009).
Adventitious shoot formation was shown to play a vital role in the development of
uniformly transformed plants from these tissues (Zhang et al., 1999). Although
adventitious regeneration is generally undesirable for clonal micropropagation, it offers
an excellent opportunity to regenerate plants from various tissues. Also the propagation
rates can be much higher than axillary shoot formation (Chun, 1993). Adventitious shoot
formation can also be used for overcoming reproductive barrier caused by sterile male/
female plants (Kantia and Kothari, 2002).
Conventional propagation techniques for woody fruit species are slow with some
inherent difficulties due to long generation cycles and high level of heterozygosity
(Sriskandarajah et al., 1994). There is a need to develop in vitro methods that could be
available to speed up the breeding process for crop improvement. Many woody plant
species resist the establishment of an efficient system for regenerating plants due to
genetically driven in vitro recalcitrance (McCown, 2000; Singh et al., 2002). However,
in vitro adventitious regeneration has been achieved from various plants of several
woody tree species (Maggon and Singh, 1996; Nagori and Purohit, 2004). It was
reported that under identical conditions the shoot regeneration percentage varied
depending on the source and type of explants used (Gentile et al., 2002; Grant and
Hammatt, 2000). A higher percentage of shoot regeneration was attained from juvenile
leaf explants as compared to adult leaves in Prunus dulcis (Miguel et al., 1996).
Regeneration has also been achieved from the leaves of apricot (Burgos and
Alburquerque, 2003), black cherry (Hammatt and Grant, 1998) and sweet cherry (Matt
and Jehle, 2005). Regeneration of adventitious shoots has been reported from immature
25
cotyledons of Peach (Yan and Zhou, 2002) and Almond (Ainsley et al., 2001). In
addition, regeneration using mature cotyledons has been reported for Peach (Pooler and
Scorza, 1995), ornamental cherries (Hokanson and Pooler, 2000) and sweet cherry (Canli
and Tian, 2008). Regeneration through adventitious shoot formation was achieved in
Feronia limonia using hypocotyl segments by Singhvi (1997).
Adventitious regeneration of Pecan has never been reported before, however, it
was reported in some members of the family Juglandaceae, e.g., Juglans nigra (Neuman
et al., 1993) and Juglans regia (Chvojka and Reslova, 1987). This phenomenon may be
of particular significance for extremely recalcitrant woody plant species such as Pecan.
Long et al., (1995) reported an unexpected observation that was the production of
adventitious shoots from the cotyledonary explants of Juglans nigra, placed on WPM
medium containing 2, 4-D and TDZ. Obeidy and Smith (1993) showed similar
adventitious buds arising from callus cultures of mature Pecan (Carya illinoensis)
embryonic tissues. Shoots were regenerated from explants placed on MS medium with
25 µM TDZ.
Later, in the experimental work of Neuman et al., (1993), no shoot organogenesis
was recorded when immature cotyledonary explants were placed on WPM medium
containing 2, 4-D and TDZ. However, Preece (unpublished data) observed shoot
organogenesis in Juglans nigra from cotyledonary explants placed on WPM medium
containing 2, 4-D and TDZ. Adventitious shoots were readily multiplied through axillary
shoot proliferation. Biotechnology utilizing adventitious regeneration may also present a
new opportunity for the improvement of woody plant species.
26
2.1.5 EFFECT OF TDZ
Thidiazuron (N-phenyl-Nَ-1, 2, 3-thiadiazol-5-yl urea; TDZ) is a synthetic
cytokinin, formerly developed by Schering and exploited as a cotton defoliant (Arndt et
al., 1976). The cytokinin-like activity of TDZ has been demonstrated by Mok et al.,
(1979) and Thomas and Katterman, (1986). TDZ is highly stable to the plants degrading
enzymes and active even at very low concentrations as compared to other synthetic
cytokinins (Mok et al., 1987). Furthermore, resistance to cytokinin oxidase contributed
to its high stability and might be a reason for its high efficacy. Its auxin-like and
cytokinin-like activity might be another possible reason for its high effectiveness (Visser
et al., 1992). TDZ also encourages the synthesis of endogenous cytokinin (Thomas and
Katterman, 1986; Victor et al., 1999) and enhances the accumulation and translocation of
auxin within the TDZ exposed tissues (Murthy et al., 1995; Hutchinson et al., 1996;
Murch and Saxena, 2001). Because of its magnificent ability to stimulate shoot
proliferation it is selected for micropropagation over a wide range of recalcitrant woody
plant species including walnut, silver maple and white ash (Huetteman and Preece,
1993). However, great care must be taken when it is employed as clonal
micropropagation. Because studies have revealed that at low levels, TDZ not only
stimulates axillary shoot proliferation but hampered shoot elongation and higher
concentration of TDZ tends to stimulate callus formation, adventitious shoos and somatic
embryos in many woody species (Huetteman and Preece, 1993). Kaveriappa et al.,
(1997) reported that TDZ at higher concentrations can cause browning and explant
necrosis, undesirable for morphogenic development.
TDZ has emerged as a highly potent regulator of morphogenetic responses in the
tissue culture of many plant species (Murthy et al., 1998; Jaiswal and Sawhney, 2006).
These responses include micropropagation (Khalafalla and Hattori, 1999; Murch et al.,
27
2000; Fratini and Ruiz, 2002), somatic embryogenesis (Saxena et al., 1992; Visser et al.,
1992; Murthy et al., 1995), breaking of bud dormancy (Wang et al., 1986), regeneration
and multiple shoot formation (Eapen et al., 1998; Li et al., 2000; Murch et al., 2000;
Chengalrayan et al., 2001; Gallo-Meagher and Green, 2002). Thus it appeared as an
efficient bioregulator of morphogenesis. The recently applied approaches to study the
morphogenic events initiated by TDZ have clearly revealed the details of a variety of
underlying mechanism (Mok and Mok, 1982; Malik and Saxena, 1992b). Some reports
indicated that TDZ might act through modulation of the endogenous plant growth
regulators, either directly or as a result of induced stress (Murthy et al., 1995;
Hutchinson and Saxena, 1996). The other possibilities included the modification in cell
membranes, energy levels, nutrient uptake, or nutrient assimilation (Chernyad'ev and
Kozlovskikh, 1990).
TDZ has also been shown to simulate shoot organogenesis from immature seeds
in several woody species such as Juglans nigra (Neuman et al., 1993) and Fraxinus
americana (Bates and Preece, 1990; Bates et al., 1992). On the contrary, Kulkarni et al.,
(2000) demonstrated that the auxin as well as cytokinins-like activities of TDZ may not
permit to induce organogenesis in internodes. Huetteman and Preece (1993) observed
that TDZ was most effective at lower concentrations (< 1µM) and induced greater
axillary proliferation but could inhibit shoot elongation. Additionally, Gairi and Rashid,
(2005) observed direct differentiation of somatic embryos on cotyledons of Azadirachta
indica on low concentrations of TDZ (0.5 µM). A higher concentration of TDZ (> 1µM),
however, could stimulate callus formation, adventitious shoots or somatic embryos. TDZ
at 10 µM regenerated adventitious shoots and somatic embryos from cotyledons of white
ash (Bates et al., 1989, 1992; Preece and Bates, 1990). Subsequent rooting of
microshoots was unaffected or slightly inhibited by prior exposure to TDZ. Undesirable
28
side effect of TDZ was that cultivars of some species occasionally formed fasciated
shoots.
In 1995, Long and coworkers initiated somatic embryos and adventitious shoots
from immature cotyledons 10-14 weeks after anthesis. They suggested that agar-
solidified WPM (Woody Plant Medium) supplemented with 0.1 µM 2, 4-D and 50 µM
TDZ and incubated in light for first 4 weeks was the best treatment for the induction of
somatic embryos and adventitious shoots from immature cotyledonary explants. Plantlets
from rooted adventitious shoots were successfully acclimatized to greenhouse
conditions.
Another important research work reported successful transfer of explants derived
from forced epicormic buds of Silver maple and Green ash (Aftab et al., 2005). In their
work, DKW medium was supplemented with 1 µM thidiazuron (TDZ) and 1 µM IBA for
axillary shoot proliferation. The results were quite satisfactory.
Based on a review of the published literature available on TDZ, it appears that an
investigation determining its role in Pecan tissue culture is lacking. This necessitates
work on this aspect that may prove beneficial for Pecan improvement.
29
CHAPTER 3
MATERIALS AND METHODS
3.1 MEDIA PREPARATION
3.1.1 PREPARATION AND STORAGE OF STOCK SOLUTIONS
Culture media, i.e., DKW medium (Driver and Kuniyuki, 1984; Annexure-I), MS
basal medium (Murashige and Skoog, 1962; Annexure-II) and WPM (Woody Plant
Medium of McCown and Llyod, 1981; Annexure-III) supplemented with various growth
regulators were tested for bud activation, callus induction and in vitro seed germination.
Stock solutions were prepared for mineral nutrients, growth hormones and vitamins
required for experimental work. Usually stock solutions were prepared in advance for the
sake of convenience and accuracy in the preparation of the above media. All solutions
were prepared using analytical grade chemicals in double distilled H2O and stored in a
refrigerator at 4oC.
For DKW (Driver and Kuniyuki, 1984) medium, stock solutions were prepared
for: Nitrates, Sulphates, Calcium, Phosphates, Iron and Organics. Detailed formulation
on DKW stock solutions is given in Annexure-I. For MS basal medium (Murashige and
Skoog, 1962) and WPM (Woody Plant Medium of McCown and Llyod, 1981), stock
solutions were prepared for: Macronutrients, Micronutrients, Iron-EDTA, Vitamins,
Myo-inositol and Growth regulators. Formulation of Murashige and Skoog (1962) stock
solutions is given in Annexure-II and for Woody Plant Medium is given in Annexure-III.
30
3.1.2 GROWTH REGULATORS
Stock solutions of growth regulators were either prepared in mM, µM or nM
concentrations and were diluted with an appropriate quantity of distilled water before
being used according to the requirement of the medium. Details for the preparation in
initial solvent and further dilution are given in Annexure-IV.
3.1.3 PREPARATION OF STOCK SOLUTIONS FOR DKW (DRIVER AND
KUNIYUKI, 1984) MEDIUM
Stock solutions of DKW are grouped together based on their compatibility. All
these stock solutions were prepared at a final concentration of 50X.
a) ORGANICS (DKW1)
All the components as given in Annexure-I under section “A” were weighed and
dissolved separately in an appropriate quantity of distilled water in 100 ml capacity
graduated beakers. Separately prepared solutions were then carefully mixed together in a
1000 ml capacity conical flask while rinsing the 100 ml capacity beakers several times
with distilled water. The contents were then transferred to a 1000 ml volumetric flask to
make the final volume with distilled water. Exactly 20 ml from this organics stock was
used for the preparation of 1 liter of DKW nutrient medium.
b) PHOSPHATES (DKW2)
Salts of phosphates as given in Annexure-I under section “B” were weighed and
dissolved separately in an appropriate quantity of distilled water. Separately prepared
solutions were mixed carefully in a 1000 ml volumetric flask so as to avoid precipitation.
Final volume was made with distilled water. It was stored in a refrigerator and 20 ml of
this phosphates stock solution was dispensed for the preparation of 1 liter of DKW
nutrient medium.
31
c) NITRATES (DKW3)
All the components of nitrates as given in Annexure-I under section “C” were
weighed and dissolved separately in an appropriate quantity of distilled water. Separately
prepared small quantities of solutions were then carefully transferred in a 1000 ml
volumetric flask to make up the final volume with distilled water. It was stored in a
refrigerator. Later 20 ml of this nitrates stock solution was used for the preparation of 1
liter of DKW nutrient medium.
d) CALCIUM STOCK (DKW4)
The salt of calcium (CaCl2) as given in Annexure-I under section “D” was
weighed and dissolved in an appropriate quantity of distilled water in a small beaker. The
solution was then transferred to a 1000 ml volumetric flask to make the final volume
with distilled water. It was stored in a refrigerator and 20 ml of this calcium stock
solution was dispensed for the preparation of 1 liter of DKW nutrient medium.
e) SULPHATES (DKW5)
All the salts of sulphates as given in Annexure-I under section “E” were weighed
and dissolved separately in an appropriate quantity of distilled water. Final volume was
made up to 1 liter as described above in section “a”. Stored in a refrigerator and
dispensed 20 ml of this sulphate stock solution for the preparation of 1 liter of DKW
nutrient medium.
f) IRON-EDTA (DKW6)
Components as given in Annexure-I under section “F” were weighed and
dissolved separately in distilled water in small beakers by using magnetic stirrer. The
solutions were then transferred to a 1000 ml capacity volumetric flask. Distilled water
was added to make up the final volume. Exactly 20 ml of this iron stock was used for the
preparation of 1 liter of DKW nutrient medium.
32
3.1.4 PREPARATION OF STOCK SOLUTIONS FOR MS (MURASHIGE AND
SKOOG, 1962) MEDIUM
a) MACRONUTRIENTS
For MS medium, the stock of macronutrients was prepared at the final
concentration of 20X as detailed in Annexure II, section “A”. All the salts were weighed
individually and dissolved separately in an appropriate quantity of distilled water.
Separately prepared solutions of different salts were mixed together in a conical flask
already containing an appropriate amount of distilled water so as to avoid precipitation.
The solution was then transferred to a 1000 ml capacity volumetric flask to make up the
final volume with distilled water. It was stored in a refrigerator. Later, pipetted 50 ml of
macronutrient stock solution for 1 liter of MS nutrient medium.
b) MICRONUTRIENTS
Stock solution of micronutrients was prepared at a concentration 100X. All the
salts of micronutrients as given in Annexure-II under section “B” were weighed
accurately and dissolved separately in approximately 100ml of distilled water. Final
volume was made up to 1 liter as described above in section “a”. It was stored in a
refrigerator and added exactly 10 ml of this stock for the preparation of 1 liter of MS
medium.
c) IRON-EDTA
Stock solution of Iron-EDTA was prepared at a concentration of 200X. The salts
as given in Annexure-II, section “C” were weighed and dissolved separately in distilled
water. The solution was then transferred to a 1000 ml volumetric flask. Distilled water
was added to make up the final volume. Iron stock was stored in an amber-colored bottle
33
in a refrigerator. For the preparation of 1 liter of MS medium, 5 ml of this stock solution
was used.
d) VITAMINS
Vitamins of MS medium were prepared at a concentration of 100X. Vitamins
were dissolved separately (as given in Annexure-II, section “D”), transferred to a 500 ml
volumetric flask and final volume was made with distilled water. It was stored in a
refrigerator. For 1 liter of MS medium, 10 ml of this vitamin stock was used.
e) MYO-INOSITOL
Stock solution of myo-inositol was prepared separately as 100X. It was prepared
by dissolving 10 g of myo-inositol in 1000 ml of distilled water and 10 ml of this stock
was taken for 1 liter MS medium.
3.1.5 PREPARATION OF STOCK SOLUTIONS FOR WPM (WOODY PLANT
MEDIUM OF McCOWN AND LLYOD, 1981)
a) MACRONUTRIENTS
Macronutrients stock for WPM was prepared at the final concentration of 20X as
detailed in Annexure-III, section “A”. All the salts of macronutrients were weighed and
dissolved separately in an appropriate quantity of distilled water in 100 ml capacity
graduated beakers. Separately prepared solutions of different salts were mixed together
in a conical flask already containing an appropriate amount of distilled water so as to
avoid precipitation. The solution was then transferred to a 1000 ml capacity volumetric
flask to make up the final volume with distilled water. It was stored in a refrigerator.
Later, pipetted out 50 ml of macronutrient stock solution for 1 liter of WPM nutrient
medium.
34
b) MICRONUTRIENTS
All the component salts of micronutrients as given in Annexure-III under section
“B” were weighed and dissolved separately in an appropriate quantity of distilled water.
The stock was prepared at a concentration of 100X. Final volume was made up to 1 liter
as described above in section a. It was stored in a refrigerator. Exactly 10 ml of this
micronutrient stock solution was used for the preparation of 1 liter of WPM medium.
c) IRON-EDTA
Stock solution of Iron-EDTA was prepared at a concentration of 20X. The salts
as given in Annexure-III, section “C” were weighed and dissolved separately in distilled
water. The solution was then transferred to a 1000 ml volumetric flask. Distilled water
was added to make up the final volume. Iron stock was stored in an amber-colored bottle
in a refrigerator. For the preparation of 1liter of WPM medium, 50 ml of this stock
solution was used.
d) VITAMINS
Vitamins of WPM medium were prepared at a concentration of 100X. Vitamins
were dissolved separately (as given in Annexure-III, section “D”), transferred to a 500
ml volumetric flask and final volume was made with distilled water. It was stored in a
refrigerator. For the preparation of 1 liter of WPM medium, 10 ml of this vitamins stock
was used.
e) MYO-INOSITOL
Stock solution of myo-inositol was prepared separately as 50X. It was prepared
by dissolving 5 g of myo-inositol in 1000 ml of distilled water and 20 ml of this stock
was taken for 1 liter WPM medium.
35
3.1.6 PREPARATION OF STOCK SOLUTIONS OF GROWTH REGULATORS
Growth regulators from various classes including auxins, i.e., Naphthalene acetic
acid (NAA), 2, 4-dichlorophenoxyacetic acid (2, 4-D), Indole-3-acetic acid (IAA),
Indole-3-butyric acid (IBA) and cytokinins, i.e., 6-benzylaminopurire (BAP) and N-
phenyl-N'-1, 2, 3-thidiazol-5-yl-urea (TDZ) were weighed and dissolved in an initial
solvent as given and explained for each in Annexure-IV. The solution was transferred to
a 50 ml volumetric flask and distilled water was added to make up the final volume.
Stock solutions of growth regulators were prepared at a concentration of 1 mM, 1 µM or
1 nM and diluted and used according to the experimental requirements. Stock solutions
of growth regulators were stored at 4°C in refrigerator till use.
3.1.7 PREPARATION OF DKW MEDIUM FROM THE STOCKS
One liter of DKW nutrient medium was prepared by taking the accurate volumes
of stock solutions given in detail in Annexure-V. Final volume was made up to one liter
by adding distilled water. The pH of the medium was adjusted to 5.7 - 5.8 by using 1.0 N
NaOH/ 1.0 N HCl. Agar (Oxoid, Hampshire, England) was added at a concentration of
7.0 g/l and the medium was heated till boiling to melt agar. The medium was then poured
in pre-sterilized culture vessels (150 × 25 mm). Culture vessels were wrapped
individually with polypropylene sheets of appropriate size and tied with rubber bands.
3.1.8 PREPARATION OF MS AND WPM MEDIUM FROM THE STOCKS
One liter of MS and WPM basal medium were prepared by taking the accurate
quantity of volumes of stock solutions given in detail in Annexure-VI and in Annexure-
VII, respectively. Final volume was made up to one liter by adding distilled water. The
pH of the medium was adjusted to 5.7 - 5.8 by using 1.0 N NaOH/ 1.0 N HCl. Agar
36
(Oxoid, Hampshire, England) was added at a concentration of 7.0 g/ l and the medium
was heated till boiling to melt agar. The medium was then poured in pre-sterilized
culture vessels (150 × 25 mm). Culture vessels were wrapped individually with
polypropylene sheets of appropriate size and tied with rubber bands.
3.2 STERILIZATION
In tissue culture procedures/ techniques, one of the vital steps is the satisfactory
sterilization of glassware, growth media, working area and surgical instruments. The
explants used must also be surface disinfested for its successful subsequent growth.
3.2.1 GLASSWARE STERILIZATION
Glassware sterilization includes the following steps:
1) All the glassware (culture-tubes, Petri-dishes, pipettes, beakers, flasks etc.)
was washed thoroughly with household detergent and given several washings
under tap water followed by a rinse with distilled water.
2) Afterwards, glassware was soaked in chromic acid over night. Chromic acid
was prepared by mixing potassium dichromate (K2Cr2O7, 10 %) and
concentrated sulphuric acid (H2SO4) in 2:1 (v/v) ratio.
3) The glassware was then washed thoroughly with running tap water by giving
several washings to remove chromic acid and sulphuric acid solution
followed by two or three rinses with distilled water.
4) Finally, the glassware was dried and sterilized in an oven at 180oC for two
hours and stored in a dust-proof cupboard till its use.
37
3.2.2 STERILIZATION OF TISSUE CULTURE MEDIA
Plant tissue culture medium, containing a high percentage of sucrose supports the
growth of microorganisms which grow faster than the cultured tissue. It is necessary to
maintain the aseptic conditions inside the culture vessels, so the culture tubes containing
different media were wrapped with polypropylene sheets of appropriate size and tied
with rubber bands. The media were sterilized by autoclaving at 15 lbs inch-2 for 15 - 20
minutes at 121oC. After autoclaving, the sterilized media were allowed to cool down at
room temperature. The sterilized media were then kept in culture room until use.
3.2.3 STERILIZATION OF GLASSHOUSE MEDIA The glasshouse media (coccopeat, sand or sawdust) were sterilized by
autoclaving at 15 lbs inch-2 for 20 - 30 minutes at 121oC and poured in flat trays for
softwood shoot forcing from large stem segments of Pecan. The sterilized sand was also
used for the rooting of softwood shoots under glasshouse conditions.
3.2.4 STERILIZATION OF WORKING AREA OF LAMINAR AIRFLOW CABINET
Aseptic transfer techniques are considered to be basic requisite for the induction
and maintenance of clean cultures free from any microbial contamination. Prior to the
inoculation or sub-culturing of explants into the culture tubes, hands, instruments and
working area of laminar airflow cabinet were cleaned and sterilized. The working area of
laminar airflow cabinet was sterilized by:
1) thoroughly scrubbing all the interior of the cabinet with 70 % ethanol.
2) irradiating with UV light for about half an hour. The UV light was
switched off at least 15 minutes before inoculation.
3) sterilizing the working bench by scrubbing with ethanol.
All the work was carried out under the gentle flow of micro-filtered air.
38
3.2.5 STERILIZATION OF SURGICAL TOOLS
All the surgical tools (scalpels, forceps, spatula, needles, scissors and blades)
used during the aseptic manipulation of explants in culture media were sterilized by
putting them in a glass-bead sterilizer (Simon Keller AG CH-3400, Switzerland) set at a
temperature of 250 ºC. All the surgical tools were kept in pre-heated glass-bead sterilizer
for couple of minutes before aseptic manipulations during working. The hot forceps and
other tools were allowed to cool down for some time to room temperature, before the
manipulation of explants into different culture media.
3.3 PLANT MATERIAL 3.3.1 SOURCE OF PLANT MATERIAL
Pecan (Carya illinoensis (Wangenh.) C. Koch) is the most important native North
American orchard species, grown primarily for commercial nut production. In Pakistan,
Pecan was first introduced in 1972 (Rehman and Jan, 1998). Many Pecan trees are
growing and fruiting in Abbotabad, Peshawar, Gilgit, Swat etc (N.W.F.P, Northern
Pakistan). Moreover, some trees had also been identified growing and fruiting in Bagh-e-
Jinnah Lahore (Punjab, Pakistan). For the present research work, Pecan trees were
selected and plant material was obtained from both of the above-mentioned areas.
3.3.2 DISINFESTATION OF PLANT MATERIAL
The surface of the plant material (buds, bark and fruits) taken from adult trees
have a variety of microbial contaminants. To get rid of these contaminants, explants were
washed under running tap water for about 15 minutes. Afterwards, the explants were
surface disinfested within a flask containing household detergent with continuous stirring
for 10 minutes and then given several rinses with distilled water. The explants were then
immersed in 15 % sodium hypochlorite (NaOCl, 3.0 % v/v) solution containing Tween
20 (0.1 % v/v) for about 10 - 20 minutes followed by 7 - 8 times rinsing with autoclaved
39
distilled water under aseptic conditions. Explants were then treated with a solution of
mercuric chloride (0.1 % w/v) for 15 minutes and rinsed 5 times with autoclaved distilled
water in laminar airflow cabinet.
3.4 EXPERIMENTAL PLAN
3.4.1 IN VITRO GERMINATION OF PECAN SEEDS
Mature seeds were collected during the mid August 2007, from trees growing in
Abbotabad. Seeds were surface disinfested in the same manner as explained in section
3.3.2. Before inoculation, hands were washed with antibacterial soap and then sprayed
with 70 % ethanol. Pecan seeds were also sprayed with 70 % ethanol, firmly held in
hands and split opened by several strokes of sterilized forcep and then placed on a clean
sterilized slab. Afterwards, one third of the fruit was excised, side lobes of the kernel
were removed carefully with sharp razor on both sides keeping the embryos intact. These
fruit pieces having intact embryos were cultured in two ways. Firstly, a small cut or
incision was made at the top tip portion to facilitate the embryo emergence/ development
and secondly, the whole tissue was cultured as such without making cuts/ incisions
(control). The explant preparatory steps for in vitro germination of seeds are explained
diagrammatically as under (Fig. 3.1 - 3.9).
Fig. 3.1 Fig. 3.2 Fig. 3.3
Fig. 3.1: A mature fruit of Pecan placed on a clean glazed ceramic slab (0.5 x).
Fig. 3.2: An enlarged view of Pecan fruit after removing the husk (1.0 x).
Fig. 3.3: Excised one third fruit of Pecan showing the excision of right cotyledon
longitudinally parallel to the embryonal axis (0.6 x).
40
Fig. 3.4 Fig. 3.5 Fig. 3.6
Fig. 3.4: A view of fruit excised parallel to the right side of embryonal axis
showing the removed cotyledon (arrow) (0.6 x).
Fig. 3.5: Another view of fruit excised parallel to the left side of embryonal axis
showing the removed cotyledon (arrow) (0.6 x).
Fig. 3.6: An enlarged longitudinal view of Pecan fruit excised on both sides
parallel to the embryonal axis (0.8 x).
Fig. 3.7 Fig. 3.8 Fig. 3.9
Fig. 3.7: A view of excised Pecan fruit showing the removal of brown testa of
fruit from the upper part of the embryonal axis (0.6 x).
Fig. 3.8: An enlarged view after the removal of brown testa on the upper part of
the embryonal axis highlighting the exposure of embryo (arrow) (0.7 x).
Fig. 3.9: A culture vessel showing the embryonal axis with the one third fruit’s
cotyledonary portion inoculated in respective medium (0.5 x).
41
Three media (DKW, MS or WPM) supplemented with six different
concentrations of BAP (0.5, 1.0, 3.0, 8.0, 12.0 or 15.0 µM) were used for in vitro
germination of Pecan seeds. These media without growth regulators were used as
control. Ten culture vessels (150 × 25 mm) were used for each combination. In total,
twenty four media involving DKW, MS or WPM containing various levels of growth
regulators (Annexure-VIII) were used. The developed seedlings were transferred after 7
days to their respective fresh nutrient media to avoid the problem of phenolic compounds
as well as to maintain the nutrient balance, i.e., inorganic salts, vitamins, sucrose and
growth regulators for further growth and proliferation/ development. Data in terms of
initiation of seed germination and percentage seed germination were recorded at day 6,
and for shoot number, root number, shoot length, root length, number of leaves and
nodes at day 25. Percentage of seed germination was calculated according to the
following equation (adapted from Maliro and Kwapata, 2000):
number of germinating seeds number of total seeds inoculated × 100 Percentage of seed germination (%) =
After reaching to an appropriate size (≥ 4 cm) the in vitro developed shoots/
plantlets were transferred to plastic pots (9.5 × 12 cm) containing perlite or vermiculite
and kept in a chamber (for one week) covered with polyethylene sheet all around in order
to maintain humidity. These pots were placed in culture room at 25 ± 2ºC and 16 h
photoperiod. Each pot was watered simultaneously according to the requirement. After 5
weeks (35 days) polyethylene sheet was removed, plantlets were transferred to plastic
pots (23 × 25 cm) containing clay-loam soil enriched with organic matter (decaying
leaves) and acclimatized initially in culture room for next 27 days. Afterwards, plants
42
were transferred to clay pots (32 × 33 cm) and maintained in glasshouse for 15 days for
further development and obtaining hardened plants of Pecan.
3.4.2 CALLUS INDUCTION AND ORGANOGENESIS
3.4.2.1 CALLUS INDUCTION FROM BARK SEGMENTS
Bark tissues from adult tree were used as an explant source. Stem segments (1.0 -
2.0 cm wide and 10.00 - 15.00 cm long) were collected during the month of August
2007, from a field-grown adult tree source. Leaves were carefully removed from the
stem segments and then further divided into smaller pieces (4.0 - 5.0 cm). Surface
disinfestation of the bark segments was carried out as explained in section 3.3.2.
Immature and mature bark tissues were removed carefully from the segments, further
divided into smaller pieces (0.5 - 1.0 cm long; 2.5 - 3.0 mm wide or 0.5 - 1.0 mm in
thickness) and with the help of forceps the segments were transferred to 150 × 25 mm
culture vessels containing 15 ml of different agar-solidified media, i.e., DKW, MS or
WPM. These media were supplemented with different concentrations of 2, 4-D and TDZ
or combination of TDZ + NAA. Ten cultures vessels (150 × 25 mm) were inoculated for
each medium tested. In total, 21 media (210 numbers of test tubes) were used and the
experiment was repeated thrice. Detailed formulation of different media tested for callus
induction and proliferation is given in Annexure-IX. Bark tissues also release phenolic
compounds in the medium, so these were transferred to their respective fresh nutrient
medium once during the experimentation after 5 days of initial culture. Data were
recorded at day 20 after first subculture. As the medium become depleted of nutrients,
therefore, in vitro agar-solidified callus cultures were shifted to their respective fresh
medium for continuous supply of nutrient and further proliferation and maintenance after
25 days of interval. Callus cultures were maintained up to 4th subculture and were hard
43
to maintain further due to sudden browning and necrosis. During different subcultures,
calluses were also transferred for plant regeneration on MS medium supplemented with
BAP (2.22 µM) or BAP + TDZ (2.22 + 0.5 or 1.0 µM). The detailed formulation of
regeneration medium is given in Annexure-X. Data in terms of callus morphology were
recorded before each subculture (at day 24) and for regeneration potential at day 25 and
35.
3.4.2.2 CALLUS INDUCTION FROM IMMATURE FRUITS
Immature fruits were collected during the month of October 2006, from an adult
field grown tree. Seeds were surface disinfested as described in section 3.3.2. Following
disinfestation, seeds were split opened by several strokes of sterilized forcep. The brown
testa was removed carefully to some extent to avoid the exudation of phenolics and
reduce lethal browning of the medium. The cotyledons were excised into smaller pieces
(0.7 - 1.3 cm long; 3.0 - 5.0 mm wide or 1.5 - 2.0 mm in thickness) and cultured on
DKW, MS or WPM media by placing one explant per 150 × 25 mm culture vessel
containing 15 ml of agar-solidified medium. Three different concentrations of 2, 4-D
(4.52, 13.57 or 22.61 µM) supplemented to these three basal media were tested. Details
of media used for callus induction and proliferation are also given in Annexure-XI. In
total, nine treatments (three 2, 4-D levels per medium) were used. Fifteen culture vessels
were used for each treatment thus making a total of 135 culture vessels. Cultures were
incubated under16 h photoperiod at 25 ± 2ºC. Data in terms of callus induction and
proliferation rate were recorded at day 30 of initial culture.
44
3.4.3 ADVENTITIOUS REGENERATION OF PECAN USING
IMMATURE COTYLEDONARY EXPLANTS
Fruits were surface disinfested as explained in section 3.3.2. Following
disinfestation, a sterilized vise was used to split open the hardened fruit. The embryo
axes were excised from fruit with great care. The isolated embryonic axes were cultured
on different media, i.e., DKW, MS or WPM supplemented with different concentrations
of BAP or TDZ (Annexure-XII) by placing in 150 × 25 mm culture vessels each
containing 15 ml of agar-solidified medium. Three culture vessels were used for each
combination of all the tested media. Cultures were incubated under 16 h photoperiod at
25 ± 2ºC. Multiple adventitious shoots were developed from the immature embryonic
axes. Data were recorded at day 23 of initial inoculation to the medium. These shoots
were transferred for rooting (without any pre-treatment) or with a pre-treatment of IBA
(1000 or 2000 ppm) to fresh MS basal medium.
3.4.4 NOVEL MICROPROPAGATION PROTOCOLS
3.4.4.1 SHOOT FORCING
Stem segments/ branches of various sizes, i.e., 20 - 25 cm long (Read and Yang,
1991) were cut from adult field-grown Pecan plant and soaked for 15 minutes in 0.78 %
NaOCl (sodium hypochlorite) solution with Tween 20 (Read and Yang, 1988, 1991;
Yang and Read, 1992, 1993). Following the bleach treatment, a fresh cut was made at the
base. The lower cut ends of the stems were immersed in containers with distilled water
containing 200 mg/l 8-hydroxyquinoline citrate (8-HQC) (Read and Yang, 1987), 30 g/l
sucrose and plant growth regulators (IBA + TDZ) at a concentration of 0.5 - 2.0 µM or
BAP (5.0 - 15 µM) in three combinations. These containers were placed in three
45
environmental conditions, i.e., culture room, glasshouse or wire house. Data were
recorded in terms of bud sprouting at day 7.
3.4.4.2 FORCING LARGE STEM SEGMENTS
An adult Pecan tree was selected and stems were cut down (140 - 150 cm long)
from juvenile tree portions (Henry and Preece, 1997a, b; Van Sambeek and Preece,
1999; Vieitez et al., 1994), further excised to large logs of 40 cm in length varying in
diameter (1.0 - 4.6 cm). These logs were forced by placing horizontally in flats (52 × 25
× 6.5 cm; L × W × H), filled with sterilized sand, coccopeat, saw dust and a control
(empty flat). Three logs per flat were taken randomly and embedded into the respective
medium and kept at 25 ± 2ºC temperature under three environmental conditions, i.e.,
culture room, wire house or glasshouse (Aftab et al., 2005). All flats contained holes at
the base for proper drainage of water. Stem segments were watered (without PGRs) daily
by hand to avoid direct water contact with emerging softwood shoots and were sprayed
(if necessary) with 0.18 % H2O2 (Aftab et al., 2005) to control microbial as well as
fungal contamination of explant.
The first run of this experiment was initiated on October 15, 2006. Softwood
shoots were harvested on December 12, 2006. The second run of the experiment was
initiated on April 14, 2007. The data pertaining to sprouting of epicormic buds from this
run were recorded on April 25, 2007. The third run of the experiment was initiated the
following year on February 24, 2008. The first harvest of softwood shoots from this
experimental run was taken on April 04, 2008 where as the second harvest was
conducted on April 20, 2008. The observations for softwood-forced shoots were
recorded per experimental unit (each 40 cm long) on % sprouting, length of softwood
46
shoots, number of forced softwood shoots, number of leaves and number of nodes on
weekly basis for both the runs.
3.4.4.2.1 ESTABLISHMENT OF SOFTWOOD SHOOTS IN DIFFERENT
ROOTING MEDIUM
Forced softwood shoots (≥ 4 cm long) were harvested with scalpels, placed in a
beaker containing distilled water so as to avoid desiccation and hand shaken gently.
Softwood shoots were given three rinses with autoclaved distilled water. To control the
transfer of possible contamination from hand great care was taken. These softwood
shoots were treated with 1000 ppm IBA, NAA or 2000 ppm IBA, NAA or in
combination of IBA + NAA (1000 + 1000 ppm) for 10 seconds, then with the help of a
sterilized forcep softwood shoots were planted in pots (9.5 × 12 cm) containing perlite,
vermiculite or sterilized sand. These pots were placed in culture room at 25 ± 2ºC
temperature and 16 h photoperiod for further growth. To maintain relative humidity,
initially pots were covered with transparent polyethylene sheet. Small pores were made
at the sides of the sheets to water the shoots daily.
3.4.5 INOCULATION OF EXPLANTS
Different explants (buds, bark and fruits) were used for different studies as
explained above. Prior to inoculation, hands and arms were washed with soap and then
sprayed with 70 % ethanol. With the help of sterilized forceps, explants were placed in
autoclaved petriplate and excised with sterilized scalpel. Culture vessels containing
different formulations of media were taken, opened near the flame of spirit lamp then
inoculated by different explants with the help of sterilized forceps. Great care must be
taken during the procedure so that forceps did not touch the agar medium. The culture
47
vessel was then wrapped again by polypropylene sheet and tied with the rubber band
after briefly heating the opening of culture vessel. The same procedure was repeated for
each culture vessel. All the culture tubes were kept in the culture room.
3.4.6 CULTURE CONDITIONS
Standard light and temperature conditions were managed in the culture room.
The cultures were placed under a 16 h photoperiod (35 µmol m-2 s-1) provided by
cool fluorescent tube lights at 25 ± 2 ºC
3.4.7 STATISTICAL DATA ANALYSIS
Analysis of variance (ANOVA) using SPSS release 12.0. software package were
applied to the data for interpretation of results. Mean values were compared and
significance of dependent variables was determined by Duncan’s or Tukey’s multiple
comparison test. Standard errors (±SE) were calculated fro each treatment.
48
CHAPTER 4
PRODUCTION OF PECAN SEEDLINGS FOLLOWING IN VITRO GERMINATION
RESULTS
4.1 IN VITRO GERMINATION OF PECAN SEEDS
During the present study, the main objective of the work was to propagate Pecan
plants using tissue culture techniques. To help achieve this objective, it was desirable to
produce a sufficient number of explants for the experimental work to proceed further.
Pecan is a recalcitrant woody plant and like many other woody trees of Juglandaceae
family, procurement of a suitable explant for micropropagation is quite a challenge.
Pecan seeds were germinated in soil under glasshouse conditions. Different media
(DKW, MS or WPM) supplemented with various levels of BAP were also tested for in
vitro seed germination. To observe the effect of medium, BAP, (or interactive effect of
medium and BAP) on in vitro seed germination, different parameters were selected
(Table. 4.1, 4.1a). The results have shown that only 13.3 % of the seeds were germinated
in glasshouse conditions, even five months after sowing of the seeds (Table. 4.1b). For in
vitro seed germination, two sets of experiments were conducted (as described in Material
and Methods, Section 3.4.1). It was observed that a better response in terms of seed
germination was obtained in seeds on which a small cut/ incision was made at tip to
facilitate embryo emergence, as compared to the control (seeds where no incision was
given) (Table. 4.1). The un-incised seeds did not show any further growth after
germination. So, further observations were continued with the incised seeds. It was
observed that maximum (96.44 %) seed germination was possible in DKW medium
49
supplemented with 4.0 µM BAP after 6.66 mean number of days of initial culture (Fig.
4.1- 4.2). Similarly, MS medium supplemented with 12.0 µM BAP was also quite
effective with 94.85 % seed germination response after 5.66 mean number of days
(Table. 4.1). It was also observed that mostly roots developed earlier and more
vigorously than the shoots (Fig. 4.4 - 4.6) while rarely both developed simultaneously
(Figs. 4.3 or 4.7). An interesting feature, i.e., formation of callus was also observed from
the point of root development and its surrounding tissues on DKW medium
supplemented with 4.0 µM BAP (Fig. 4.8 - 4.9). A significant variation was observed in
seed germination percentage, however, no significant effect of medium, BAP or in
combination (Medium + BAP) was recorded on in vitro Pecan seed germination in terms
of germination period (days) (Table. 4.1, 4.1a).
50
TABLE. 4.1: EFFECT OF DKW, MS AND WPM MEDIUM SUPPLEMENTED WITH VARIOUS LEVELS OF BAP ON IN VITRO SEED (INCISED)
GERMINATION AND OTHER MORPHOLOGICAL PARAMETERS OF PECAN AT 25 TH DAY OF INITIAL CULTURE
Medium BAP
(µM) Germination
Period (days)
Seed Germination
(%)
Number of
Shoots
Shoot Length
(cm)
Number of Leaves
Number of Nodes
Number of
Roots
Root Length
(cm)
0.0 6.00 (13.55)*
84.81 bcd
(11.63)j1.16 d 3.12 cde 8.52 bcde 1 a 5.92 de 3.12 cde
1.0 7.33 (9.44)
94.62 ab
(22.99)d1.23 d 3.39 cde 5.33 b 7.85 bcdef 1 a 3.36 efg
4.0 6.66 (8.77)
96.44 a
(30.18)b5.68 a 4.03 abc 10.42 a 18.02 a 1 a 13.86 a
8.0 6.66 (9.22)
90.36 abc
(42.19)a3.46 bc 2.75 efgh 10.21 a 16.64 a 1 a 1.73 g
12.0 6.66 (9.33)
91.88 abc
(15.21)fgh1.13 d 4.36 ab 4.52 bcd 4.86 cdef 1 a 8.76 bc
DKW
15.0 6.33(8.66)
85.22 bcd
(18.52)ef1.8 cd 1.96 gh 2.73 cde 4.13 f 1 a 5.1def
0.0 6.00 (11.44)
89.55 abc
(13.29)hij1.73 cd 2.06 fgh 2.89 cde 10.61 b 1 a 4.4 defg
1.0 6.66 (8.22)
92.18 abc
(20.11)de1.16 d 2.76 efgh 1.56 e 6.99 bcdef 1 a 5.22 def
4.0 6.66 (8.77)
94.85 ab
(21.11)de1.56 d 3.1 cdef 4.70 bc 7.24 bcdef 1 a 2.98 efg
8.0 6.33 (9.11)
93.33 ab
(30.29)b1.63 d 3.93 abcd 2.5 de 6.88 bcdef 1 a 1.88 g
12.0 5.66 (8.88)
82.17 cde
(22.77)d 2.1 bcd 4.06 abc 4.31 bcd 9.33 bcd 1.16 a 3.03 efg
MS
15.0 6.33 (8.55)
90.18 abc
(26.70)c1.08 d 1.63 h 1.73 e 4.9 ef 1 a 2.76 fg
0.0 5.00 (13.22)
79.90 cde
(10.36)j1.59 d 2.23 fgh 2.85 cde 7.77 bcdef 1.16 a 5.3 def
1.0 6.33(11.66)
72.59 e
(15.22)fgh1.34 d 3.64 bcde 1.85 e 9.47 bc 1 a 4.55 defg
4.0 6.33 (9.33)
77.88 de
(14.36)ghi3.64 b 2.91 efgh 10.29 a 16.48 a 1 a 11.13 b
8.0 6.66 (8.33)
71.55 ef
(13.00) hij1.3 d 3.95 abcd 4.57 bcd 6.47 bcdef 1 a 7.10 cd
12.0 6.00 (9.11)
76.73 de
(17.65)efg1.23 d 4.88 a 2.83 cde 5.06 def 1a 2.73 fg
WPM
15.0 5.66(9.99)
76.73 de
(20.19)de1.16 d 1.93 gh 4.46 bcd 4.06 f 1 a 1.83 g
Data presented here are the means of 3 values per BAP treatments. Different letters within a specific column represent significant difference at P<0.05 according to Duncan’s Multiple Range Test.
* The values in parentheses represent the data for the seeds given no incision treatment.
The data for morphological features were recorded at day 25 except for seed germination period (days) or seed germination percentage
51
TABLE. 4.1a: ANALYSIS OF VARIANCE OF DIFFERENT PARAMETERS FOR IN VITRO SEED (INCISED) GERMINATION OF PECAN AT 25 TH DAY OF INITIAL
CULTURE
Mean square Source of
variation
df Germination
Period (days)
Seed Germination
(%)
Number of
Shoots
Shoot Length
(cm)
Number of Leaves
Number of Nodes
Number of
Roots
Root Length
(cm) Medium (A)
2 0.222NS 131.488** 4.641NS 0.567NS 41.922** 26.251** 4.630NS 44.240**
BAP (B) 5 0.922NS 94.784** 3.936NS 4.656** 43.686** 92.762** 7.407NS 48.125**A × B 10 0.844NS 101.317** 2.209NS 2.270** 11.399** 40.715** 1.019NS 23.244**
NS Non-Significant ** reflects significance at P<0.05 value according to F test with df mentioned against each.
TABLE. 4.1b: GLASSHOUSE AND IN VITRO-GERMINATION OF PECAN SEEDS
Type of culture Plant material Number of explants sown/ cultured
Germination (%)***
Glasshouse Seed 75 13.3 In vitro Seed 440 86.942
*** The percentage presented here is a compilation of data obtained in the three replicates of this experiment. Data were recorded on the 6th day of incubation for in vitro and at 47th day for glasshouse experiments.
52
Figure 4.1 - 4.10: In vitro-germinating Pecan seeds on DKW, MS or WPM basal media placed under 16 h photoperiod at 25 ± °C
Root
Shoot
Fig. 4.1 Fig. 4.2 Fig. 4.3
Fig. 4.1: Root initiation (arrow) on WPM medium at day 6 of initial culture, showing
also the cotyledonary portion of cultured fruit (right bracket) (1.6 x).
Fig. 4.2: Elongation of root (in the direction of curved arrow) on WPM medium at day
8 of initial culture (1.3 x).
Fig. 4.3: Simultaneous development of shoot and root on WPM medium supplemented
with 1µM BAP at day 6 of initial culture (1.3 x).
Fig. 4.4 Fig. 4.5 Fig. 4.6
Fig. 4.4: Elongation of root (curved arrow) on DKW medium at day 8 of initial culture
(1.0 x)
Fig. 4.5: Elongation and further development of root at day 9 of initial culture. An arc
shows the cotyledonary portion of fruit cultured on DKW medium (1.3 x).
Fig. 4.6: Another view (almost at right angel to the view in Fig. 4.5 showing the
development of root (2.0 x).
53
Shoot Root
Callus
Callus
Shoot
Fig. 4.7 Fig. 4.8 Fig. 4.9 Fig. 4.10
Fig. 4.7: Simultaneous development of both shoot and root (double headed arrow) on
MS basal medium at day 9 (1.3 x).
Fig. 4.8: Development of shoot (shorter arrow) and root (violet arrow) with the
formation of callus (a curved line over green area) from the fruit portion
adjoining the embryonal axis on DKW medium supplemented with 4 µM
BAP at day 15 (Front view, 1.3 x).
Fig. 4.9: An enlarged view of Fig. 4.8 highlighting the formation of callus (arrows)
from the fruit portions adjoining the embryonal axes on DKW medium
supplemented with 4µM BAP at day 15 (2.5 x).
Fig. 4.10: An opposite view of Fig. 4.8 showing the development of root (arrow) at
1.3 x.
54
During the culture period, multiple shoots were also observed from intact nodal
regions of developing seedlings. However, the maximum number of multiple shoots
(5.68) were observed in DKW medium (Fig. 4.11 - 4.13) followed by those in WPM
medium (3.64) (Fig. 4.14 - 4.16) both supplemented with 4.0 µM BAP at day 25 (Table.
4.1). In DKW medium supplemented with 8 µM BAP, a bunch of multiple shoots
developed from a single initial point (Fig. 4.17) while in MS medium supplemented with
12 µM BAP, two shoots were produced per culture vessel (Fig. 4.18). However,
maximum shoot length (4.88 cm) was observed on WPM medium supplemented with 12
µM BAP (Fig. 4.19) as compared to an average (4.36 cm) on DKW basal medium. DKW
medium supplemented with 4.0 µM BAP showing highest mean number of shoots
(5.68), leaves (10.42), nodes (18.02) or root length (13.86 cm) proved to be the best
combination in terms of multiple shoot development (Table. 4.1). WPM medium
supplemented with 4.0 µM BAP resulted in 10.29 mean numbers of leaves at day 25 of
initial culture (Table. 4.1). There was no significant effect of medium, BAP or their
combination on the data pertaining to mean number of shoots and roots. Furthermore,
medium (A) and BAP (B) alone or their interaction (A × B) had significant effect on
mean number of leaves, nodes and root length at the P<0.05 level (Table 4.1a).
Maximum number of nodes (18.02) were observed in DKW medium supplemented with
4 µM BAP (Fig. 4.11-12) with maximum root length of 13.86 cm (Fig. 4.20) followed by
11.13 cm on WPM medium (Fig. 4.14) both supplemented with 4.0 µM BAP (Table
4.1). A more pronounced and significant effect of medium, BAP or both was observed in
terms of root development (Fig. 4.13- 4.16; 4.20- 4.22). During the experiments, an
exciting feature, i.e., formation of root nodules were also observed on DKW medium
supplemented with 4.0 µM BAP (Fig. 4.20 - 4.22). These nodules were globular in shape
55
and pale-yellowish in appearance. These were seen developing on the secondary and
tertiary roots. Such structures have never been seen in any other culture.
Figure 4.11 - 4.23: Multiple shoots raised from in vitro-germinating Pecan seedlings
on DKW, MS or WPM medium supplemented with various levels of BAP at day 25 under 16 h photoperiod at 25 ± 2 °C
Multiple shoots
Fig. 4.11 Fig. 4.12 Fig. 4.13
Fig. 4.11: Multiple shoot formation (curve) on DKW medium supplemented with 4 µM
BAP (0.5 x). Fig. 4.12: An enlarged and opposite view of multiple shoots (brace) formed on DKW
medium supplemented with 4µM BAP. An arrow indicates the remaining fruit
portion
(1.3 x).
Fig. 4.13: Multiple shoot formation (arrows) on DKW medium supplemented with 8 µM
BAP (0.7 x).
Fig. 4.14 Fig. 4.15 Fig. 4.16
56
Fig. 4.14: Multiple shoot (small arrows) formation on WPM medium supplemented with
4 µM BAP, a bigger arrow indicating the direction and development of root
(1.0 x).
Fig. 4.15: An opposite view of Fig. 4.14 (curved arrow) indicating the direction of
further development of root from in vitro germinating seedling of Pecan (1.0
x).
Fig. 4.16: A left side view of Fig. 4.15 showing the vigorous root development (a longer
arrow). A short arrow indicates secondary root development on WPM
medium supplemented with 4 µM BAP (1.0 x).
Remaining fruit part
Fig. 4.17 Fig. 4.18 Fig. 4.19
Fig. 4.17: A bunch of multiple shoots arising from single initial point on DKW medium
supplemented with 8 µM BAP also showing the remaining fruit portion (0.9
x).
Fig. 4.18: Two shoots (double headed arrow) developed on MS medium supplemented
with 12 µM BAP, also showing the cotyledonary portion of fruit (two single
arrows) (1.3 x).
Fig. 4.19: In vitro-germinating Pecan seedling showing maximum shoot length with
upward root highlighting two nodular structures (arrow) at the root tip formed
on WPM medium supplemented with 12 µM BAP (0.7 x).
57
Fig. 4.20 Fig. 4.21 Fig. 4.22
Fig. 4.20: Development of primary root with the formation of secondary roots (right
bracket, arrows) on DKW medium supplemented with 4 µM BAP at day 25
(front view to the embryonal axis) (0.3 x).
Fig. 4.21: An opposite and full view of Fig. 4.20 showing the formation of multiple
shoots (arrows) and nodular structures (left bracket) on secondary roots at day
25 of initial culture (0.5 x).
Fig. 4.22: An enlarged view of Fig. 4.21 (lower half) showing the formation of nodular
structures (arrows) on secondary roots at day 25 of initial culture (1.0 x).
58
4.2 ROOTING OF IN VITRO MULTIPLE SHOOTS DEVELOPED FROM IN VITRO-GROWN SEEDS OF PECAN
At day 35 of the culture initiation, the developed multiple shoots from in vitro
germinating seedlings acquired sufficient length (3 - 4 cm) and were ready for transfer to
the rooting media. Two media (DKW or MS) with different combinations of growth
regulator (IAA, IBA or NAA) were tested for rooting. The results of the experiment for
rooting of multiple shoots are given in Table 4.2. The data reveal that none of the shoots
rooted in MS medium supplemented with 8.0 µM NAA or in DKW medium
supplemented with 2.0 µM IBA (Table. 4.2). However, the shoots transferred to MS
medium with 8.0 µM IBA or a combination of 6.0 µM IAA + 6.0 µM IBA resulted in the
formation of callus at the base of shoots rather than the development of roots (Fig. 4.23 -
4.24). Best rooting of the multiple shoots (Fig. 4.25) was obtained on MS medium
supplemented with 4.0 µM IBA + 4.0 µM NAA. On this medium, callus formation at the
shoot base with 88.46 % rooting was observed at day 24 with mean number of 1.36 roots
per culture vessel having an average root length of 2.76 cm. In 15.38 % culture vessels,
MS medium supplemented with 4.0 µM IBA + 4.0 µM NAA favored formation of two
roots as well as callus (Fig. 4.26 - 4.28), whereas only callus formation was observed in
7.69 % culture vessels (Fig. 4.29 - 4.30). Rarely, in the same medium stated above
healthy roots were developed but shoot development ceased and resulted in shoot
necrosis/ death (Fig. 4.31 - 4.33). An average (77.66 %) rooting of multiple shoots was
observed on MS medium supplemented with 4.0 µM NAA with 1.10 mean number of
roots having 2.4 cm mean root length (Table. 4.2). DKW medium supplemented with 4.0
µM NAA, however, favored only root initiation but no further development was
recorded (Fig. 4.34 - 4.35).
59
Figure 4.23 - 4.35: Rooting of in vitro multiple shoots developed from in vitro-grown seeds of Pecan
Fig. 4.23 Fig. 4.24 Fig. 4.25 Fig. 4.26 Fig. 4.27
Fig. 4.23: Multiple shoots transferred to MS medium supplemented with 8 µM NAA
showing the formation of light-green friable callus with white patches at the
shoot base (arrow) at day 15 of transfer to rooting medium (1.0 x).
Fig. 4.24: Shoot transferred to DKW medium supplemented with 2 µM IBA showing the
formation of friable, transparent and brown callus at the shoot base (arrows) at
day15 of transfer to rooting medium (1.0 x).
Fig. 4.25: Shoot transferred to MS medium supplemented with 4 µM IBA + 4 µM NAA
showing the formation of compact, yellowish-brown callus at the shoot base
and developed root (arrow) after 35 days of transfer to rooting medium (0.8 x).
Fig. 4.26: Formation of two roots (arrows) with brownish, compact callus at the shoot
base on MS medium supplemented with 4 µM NAA at day 32 of transfer to
rooting medium (0.9 x).
Fig. 4.27: An opposite view of Fig. 4.26 highlighting the formation of callus, multiple
shoots (arrows) and roots (0.9 x).
C
MS Callus
Callus
R R
Fig. 4.28 Fig. 4.29 Fig. 4.30 Fig. 4.31
60
Fig. 4.28: Formation of two roots (two combined arrows) with yellowish brown, compact
callus at shoot base on MS medium supplemented with 4 µM IBA + 4 µM
NAA at day 30 of transfer to rooting medium (1.0 x).
Fig. 4.29: Formation of compact, transparent, watery callus (arrows) at shoot base with no
root on MS medium supplemented with 4 µM IBA + 4 µM NAA at day 26 of
transfer to rooting medium (1.0 x).
Fig. 4.30: An enlarged and opposite view of Fig. 4.29 showing the formation of creamy-
white callus (arrows) at base of multiple shoots (MS-arrows) (1.3 x).
Fig. 4.31: Root induction (arrow) with transparent, yellowish-brown, compact callus
formed (double arrows) on MS medium supplemented with 4 µM IBA + 4 µM
NAA at day 25 (1.2 x).
R
C
S R
Callus
Fig. 4.32 Fig. 4.33 Fig. 4.34 Fig. 4.35
Fig. 4.32: Vigorous callus growth with development of root (longer arrow) on MS
medium supplemented with 4 µM IBA + 4 µM NAA at day 32 showing shoot
necrosis and ultimately shoot death (shorter arrow) (0.9 x).
Fig. 4.33: Browning of callus, swelling and browning of the root at day 37 of transfer to
rooting medium (0.9 x).
Fig. 4.34: Multiple shoots with the formation of compact, brown callus showing root
initiation (arrow) on MS medium supplemented with 4 µM NAA at day 25 of
transfer to rooting medium (0.7 x).
Fig. 4.35: An enlarged view of Fig. 4.34 highlighting the root induction (arrow) and
necrosis of shoots showing no further growth of root at day 35 (0.9 x).
61
TABLE: 4.2 EFFECTS OF DKW AND MS MEDIUM WITH DIFFERENT LEVELS OF IAA, IBA OR NAA ON ROOTING IN PECAN
Growth Regulators (µM)
Medium IAA IBA NAA
Root
induction
(days) A
Number of
roots A
Root
length
(cm) A
Rooting
(%) A
- - 4.0 25.33 a 1.03 b 0.96 c 32.25 d
- - 8.0 23.66 a 1.06 b 1.56 b 48.92 cDKW
- 2.0 - NRB
- 4.0 4.0 24.66 a 1.36 a 2.76 a 88.46 a
- - 8.0 NRB
- 8.0 - Callus formation at shoot base
- - 4.0 24.33 a 1.10 b 2.4 a 77.66 bMS
6.0 6.0 - Callus formation at shoot base
A Data presented here are the means of 3 values per treatment B NR represents that no roots were developed. Different letters within a specific column represent significant difference at P<0.05 according to Duncan’s Multiple Range Test.
62
4.3 HARDENING AND ACCLIMATIZATION OF IN VITRO-GROWN
PLANTS OF PECAN
At 40th day of the initial cultures, the developed plants/ seedlings acquired
sufficient length (4 - 5 cm) and were ready to be transferred for hardening. The usual
protocol employed for hardening of the well-established in vitro-grown Pecan
plants/seedlings was described in Materials and Methods, section 3.4.1. These in vitro-
grown Pecan plants were hardened successfully in perlite or vermiculite medium (Fig.
4.36 - 4.45). Some of the hardened plants showed necrosis at the leaf tips (Fig. 4.41,
4.42) that resulted in complete death of the seedlings after 27 days (Fig. 4.43). More than
85 % of in vitro-grown Pecan plants were acclimatized successfully to the glasshouse
conditions where they are flourishing very well. A figurative description of various steps
involving the hardening of in vitro-raised plantlets of Pecan in perlite or vermiculite
medium is depicted in Fig. 4.36 - 4.45.
Fig. 4.36 - 4.45: Hardening and acclimatization of in vitro-grown plants of Pecan kept in culture room for 30 days at 25 ± 2 °C
Perlite
Fig. 4.36 Fig. 4.37
Fig. 4.36: In vitro-grown Pecan seedling transferred in perlite showing the remaining
cotyledonary part (arrow) of fruit at day 1 (0.5 x).
Fig. 4.37: The Pecan plantlet in perlite at 7th day (0.3 x).
63
Vermiculite
Fig. 4.38 Fig. 4.39
Fig. 4.38: In vitro-grown Pecan seedling being hardened in vermiculite showing the
remaining cotyledonary part (arrow) of fruit at day 1 (0.8 x).
Fig. 4.39: The Pecan seedling at 7th day of transfer in vermiculite (0.3 x).
Fig. 4.40
Fig. 4.40: The in vitro-grown Pecan seedlings in perlite and vermiculite kept in an
artificially constructed chamber (arrow showing polyethylene sheet) for
retention of humidity (0.1 x).
64
Fig. 4.41 Fig. 4.42 Fig. 4.43
Fig. 4.44 Fig. 4.45
Fig. 4.41: Browning of the leaves has just begun from the tip (arrows) in vermiculite at
day 11 (1.0 x).
Fig. 4.42: Browning of the leaves extended towards the leaf base in vermiculite at day
19 (1.3 x).
Fig. 4.43: The death of Pecan plantlet at day 27 (1.3 x).
Fig. 4.44: In vitro-grown Pecan plantlets in perlite and vermiculite kept in an artificially
constructed chamber (polyethylene sheet) at day 15 (0.2 x).
Fig. 4.45: In vitro-grown Pecan plantlets at day 30 (0.2 x).
65
Fig. 4.46 - 4.49: Acclimatized in vitro-grown plants of Pecan under glasshouse conditions
Fig. 4.46 Fig. 4.47 Fig. 4.48
Fig. 4.49
Fig. 4.46: Pecan plant kept in culture room for 15 days at 25 ± 2 °C after acclimatization
(0.3 x).
Fig. 4.47: Acclimatized Pecan plant under glasshouse conditions at 45th day under
natural light conditions 25 ± 2 °C (0.25 x).
Fig. 4.48: Acclimatized Pecan plant in glasshouse at 65th day (0.25 x).
Fig. 4.49: Acclimatized well developed in vitro-raised plants of Pecan ready for their
transfer to field conditions (0.15 x).
66
DISCUSSION
Pecan (Carya illinoensis) is an excellent multipurpose, hard-wood tree species of
commercial importance mostly valued for its nut crop and furniture grade wood. Pecan is
usually propagated by seed. Grafting and budding on seedling rootstocks are the other
propagation methods of Pecan under nursery conditions (Smith et al., 1974; Menary et
al., 1975). Pecan propagation either by budding or grafting is considered difficult yet is
used in some cases (Young and Young, 1992). On the other hand, these methods suffer
disadvantages such as considerable time and poor transplanting survival of the plants.
Further, these methods were also not sufficiently reliable or adequate to meet the
growing demand of Pecan nuts and their products. Since conventional propagation
methods such as using seeds, budding and grafting have not succeeded in producing
large quantities of Pecan propagules, it was necessary to investigate alternative strategies
for successful propagation. In addition, for a number of other reasons including limited
availability of seed material and need for rapid propagation seeds may be germinated
under in vitro conditions.
The application of tissue culture methods offers great potential for propagation
and improvement of Pecan as described for other woody plants (Litz, 1984). Previous
studies with Pecan tissue culture have shown that it is difficult to propagate this
recalcitrant tree species through in vitro techniques (Wood, 1982). On the other hand,
several workers have reported successful attempts on various aspects of research on
Pecan (Hansen and Lazarte, 1984; Burns and Wetzstein, 1997; Grauke et al., 2003;
Beedanagari et al., 2005). In vitro culture is an efficient method for vegetative
propagation as well as ex-situ conservation of plant diversity (Krogstrup et al., 1992;
Fay, 1994). Hence, use of in vitro protocols has been anticipated as a successful
approach for ex-situ conservation and re-introduction of endangered plant species
67
(Stenberg and Kane, 1998; Decruse et al., 2003; Sarasan et al., 2006). Several authors
have noted certain advantages of using seeds that is, intact seedlings as primary explants
(Malik and Saxena, 1992a, 1992b, 1992c; Victor et al., 1999). In in vitro studies, seeds
are preferred as starting material for establishing cultures (Benson, 2000), as they are the
representative of the genetic structure of the target population to be conserved (Alves et
al., 2006). Additionally, propagation from seed is desirable because it is logically simple
(Yildrim et al., 2007). The in vitro germination of seeds allows a yield of a large number
of aseptic plants to be inoculated in tissue culture (Mercier and Kerbauy, 1997). Maliro
and Kwapata (2000) demonstrated that in vitro conditions achieve high germination
percentages and provide aseptic and juvenile plants for rapid micropropagation.
Furthermore, plants regenerated from seeds have a broader genetic background than
those developed by clonal propagation methods (Munoz and Jimenez, 2008). Therefore,
in vitro seed germination holds great promise in overcoming the difficulties encountered
in propagation of plants through conventional methods. In the present study, mature
Pecan seeds were used for in vitro germination and micropropagation. In vitro seed
germination was undertaken as an alternative propagation technique other than
conventional one to obtain physiologically active and clean cultures for their proposed
utilization during the present work. Generally, propagation in Pecan (or genus Carya) is
achieved through seeds and other propagation methods as mentioned above but in
contemporary literature in vitro seed germination involving growth regulators is scanty.
During the present study, it was observed that under glasshouse conditions, the
percentage germination (13.3 %) response of Pecan seeds was very low. In contrast to
our study, Yildirim et al., (2007) obtained 50 % seed germination in P. armeniaca under
glasshouse conditions indicating recalcitrant nature of Pecan. Germination of Pecan seed
was hampered due to hard seed coat. The husks physically impede the elongation of the
68
radicle. Generally, hickories exhibit embryo dormancy, but previous work with Pecan
suggests that mechanical restriction by the hard shell is the reason for the delayed
germination in this species (Van Staden and Dimalla, 1976). Studies have also shown
that the hard seed coat renders the seeds impermeable to water and oxygen needed for
germination process (Baskin and Baskin, 1998). Similarly Maliro and Kwapata (2000),
in a study involving Uapaca kirkiana, demonstrated that the presence of hard outer seed
coat layers delays seed germination due to impermeability and restriction of radicle
emergence. Higher percentage seed germination was achieved when outer and inner seed
coat layers were removed completely (Prins and Maghembe, 1994). Due to the poor
results obtained under glasshouse conditions in this study, in vitro germination of Pecan
seeds was carried out after carefully removing the outer hard husk of seeds. During the
present work, effect of different media (DKW, MS or WPM) was investigated for in
vitro germination of Pecan seeds. The results of present investigation revealed that media
plays a significant role in in vitro seed germination. In one experimental set where seeds
were not given incision treatment, 13.29 % seeds germinated on MS basal medium
followed by DKW (11.63 %). WPM medium favored only 10.36 % seed germination. In
another set of experiment similar results were obtained as MS basal medium favored
highest (89.55 %) germination followed by DKW medium (84.81 %) in seeds given
incision treatment. Once again WPM medium yielded the lowest (79.90 %) germination
response. These results are in strong harmony with the previous studies that in vitro
germination of most plant seeds was achieved by use of basal salts medium (Kurt and
Erdag, 2009). Additionally, Murashige and Skoog (MS) formulation was the most
commonly used medium in plant tissue culture experiments (Molia, 2000). On the other
hand, in a study involving Centaurea zeybekii, Kurt and Erdag, (2009) obtained highest
germination (80 %) in distilled water containing various vitamins and 1mg/ l GA3 rather
69
than the other media (B5, MS and White’s media). Nonetheless, they found that MS
medium yielded the lowest germination response. The results of present research work
proved MS basal medium to be the most suitable medium for in vitro seed germination
of Pecan. Hopkins and Huner, (2004) also demonstrated that the presence of large
quantities of stored carbon, mineral elements and hormones in cotyledons may be
responsible for the occurrence of germination in PGR-free medium that also support the
growth and development of seedling. Furthermore, mineral demand during the process of
germination depends upon the species and is probably related to the amount of reserves
in the seed, genotypes, seed age, size and growth factors might be affecting the
germination of seeds in vitro (Padilla and Encina, 2003). The results of present work also
revealed that incision on the seeds may perhaps facilitate the embryo emergence that
accounts for the higher germination percentage in incised seeds. During the present
studies, effects of various levels of benzylaminopurine (BAP) were also investigated on
in vitro seed germination. Although germination occurred without the addition of
cytokinin to the medium, i.e., basal medium, but the rate of seed germination was
significantly improved on medium containing BAP. The present investigation
demonstrates that in non-incised seeds highest (42.19 %) germination was achieved with
8.0 µM BAP followed by 31.18 % with 4.0 µM BAP. The addition of 8.0 µM BAP
favored 30.29 % seed germination. While in incised seeds highest germination (96.44 %)
was observed on DKW medium with 4.0 µM BAP, followed by MS medium containing
4.0 µM BAP (94.85 %). WPM medium yielded the highest (77.88 %) germination
response at 4.0 µM BAP. This effectiveness of various levels of BAP in basal nutrient
medium was confirmed by Yildirim et al., (2007). They obtained 75 % seed germination
in Prunus armeniaca, with 4.43 µM BAP. The exact function of cytokinin in
germination is unknown but there is evidence that in seeds with high levels of storage
70
lipids such as Pecan nuts, cytokinins play an important role in lipid mobilization
(Dimalla and Van Staden, 1977). The requirement for cytokinin in germination media
may thus be related to utilization of lipids (De Pauw et al., 1994). It has also been shown
that if storage lipids can not be utilized, germination will not continue (Manning and Van
Staden, 1987). The promotive effect of cytokinin is also related to the alleviation of
internal stress factors (Nikolic et al., 2006). Chiwocha et al., (2005) also demonstrated
the participation of cytokinins in the development and metabolism of all phases of
seedling growth. In the view of the results of present research work, it was established
that amongst all the media and tested levels of BAP, DKW medium supplemented with
BAP at 4.0 µM proved to be the best combination tested.
The present investigation also highlighted different morphogenic responses of the
cultured explants. The addition of various levels of BAP has stimulated multiple shoot
formation from in vitro germinating seedlings. In most instances, multiple shoots
developed as a result of proliferations of pre-existing meristems in cotyledonary nodes,
shoot tips and epicotyl (Shri and Davis, 1992; Subhadra et al., 1998) were often not
reproducible. During this study, highest number of multiple shoots (5.68) was observed
on DKW medium supplemented with 4.0 µM BAP followed by WPM medium
supplemented with 4.0 µM BAP (3.64). Though more multiple shoots were induced at
lower concentration of BAP (4.0 or 8.0 µM), the number of elongated shoots of size >
4.0 cm was higher at higher concentration of BAP (12 µM). It was also observed that
multiple shoots proliferated well upon transfer to their respective media. Jayanand
(2003) also observed high proliferation of shoots when subcultured on the same medium.
During the present work, BAP was found to be an important factor in in vitro seed
germination as well as in the development of multiple shoots. In previous reports, BAP
was used as principal hormone for the induction of multiple shoot buds. During the
71
present research work, it was also observed that in 85 % of the culture vessels, roots
developed earlier than the shoots. Embryo cultures lacking roots with delayed growth of
shoots have also been reported for mature and non-mature embryos (Fregene et al.,
1999). A most interesting phenomenon, i.e., the formation of green, compact callus at the
adjoining cotyledonary portions or from the point of shoot origin was also observed on
DKW medium supplemented with 4.0 µM BAP. Another remarkable feature, i.e.,
swelling of the secondary and tertiary roots was also observed at some points with the
vigorous root development on the same medium. This might be due to lesser availability
of space in the culture vessel for vigorously-produced primary roots.
Previous studies on Pecan showed that rooting with any of the media
combination resulted in a very low frequency of root formation and most of the non-
rooted shoots died. Similarly, during the present investigation, no root induction of
shoots was observed upon transfer to MS medium containing NAA or DKW medium
supplemented with IBA. MS formulation with IBA (6.0 or 8.0 µM) resulted in the
formation of profused callus at the shoot base. In several plant species, IBA was reported
to be the most favorable auxin for root formation (Yadav et al., 1990; Pevalok-Kozlina et
al., 1997; Thomas, 2007). Previously, Wood (1982) also reported proliferation and
elongation of Pecan shoots but unable to subculture shoots and induce rooting. However,
Hansen and Lazarte (1984) developed highest number of roots on WPM with IBA (1.0 or
3.0 mg/ liter) treatment. MS medium with a combination of IBA (4.0 µM) and NAA (4.0
µM) promoted maximum (88.46 %) root induction followed by MS medium containing
NAA resulted in the formation of callus at the shoot base. This study also revealed that
the mean number of roots and root length was more in MS medium supplemented with
IBA and NAA as compared to the medium in which only NAA was added. This showed
that a combination of IBA and NAA (4.0 + 4.0 µM) was the best in terms of root
72
induction in Pecan. The present work also revealed that the Pecan seedlings developed
under in vitro conditions were successfully acclimatized to perlite or vermiculite and
more than 85 % of the plants were transferred to soil under glasshouse conditions. These
well growing plants were then successfully established in field.
In conclusion, results of the present research work demonstrated three important
aspects. Firstly, in vitro conditions favored seed germination in Pecan more than the
glasshouse conditions. Secondly, DKW medium with BAP at 4.0 µM was found to be
the best treatment for in vitro seed germination of Pecan. Finally, due to several
limitations in conventional breeding procedures, this protocol may help to produce a
sufficient Pecan stock and may also serve as an alternative or complement successful
pathway for the propagation of this recalcitrant tree to the existing germination
techniques. Thus, this procedure not only enables production of large number of aseptic
seedlings in short duration but also plays an important role in multiplication,
establishment and improvement of Pecan.
73
CHAPTER 5
NOVEL MACRO/MICRO-PROPAGATION METHODS
RESULTS Forcing shoot tips and/ or epicormic latent buds from the large stem segments of
older trees is an alternative approach for clonal propagation of trees. The shoot tips or
large stem segments were cut from the more juvenile portion during the dormant season
providing the use of plant material for a longer period of the year with reference to
propagation. The present work has been conducted with an aim to produce a sufficient
amount of Pecan stock. Due to several limitations in conventional breeding procedures, it
also aims to develop certain newer means of multiplication and establishment for this
recalcitrant tree species (Fig. 5.1).
5.1 SHOOT TIP FORCING
The stem segments from adult Pecan tree were harvested during the dormant season
and the lower cut ends of the stems were immersed in three different forcing solutions
under three different environmental conditions, i.e., culture room, glasshouse or wire
house/ natural (Fig. 5.2). The detailed formulations of these media were explained in
Materials and Methods, section 3.4.5.1.
The stem segments placed in forcing solutions under culture room conditions did not
show any response in terms of bud break (Fig. 5.2), however, contamination at shoot
base was observed in all the tested forcing solutions in culture room at 25 ± 2 ºC (Fig.
5.3 - 5.6).
74
BA
Fig. 5.1: A mature Pecan tree from Lahore, Pakistan. A) photographed in May, 2008
B) the same tree as in A during dormant season (December, 2008).
0102030405060708090
100
IBA
+TD
Z+B
AP
(0.5
+0.5
+5.0
)IB
A+T
DZ+
BA
P(1
.0+1
.0+1
0.0)
IBA
+TD
Z+B
AP
(2.0
+2.0
+15.
0)IB
A+T
DZ+
BA
P(0
.5+0
.5+5
.0)
IBA
+TD
Z+B
AP
(1.0
+1.0
+10.
0)IB
A+T
DZ+
BA
P(2
.0+2
.0+1
5.0)
IBA
+TD
Z+B
AP
(0.5
+0.5
+0.5
)IB
A+T
DZ+
BA
P(1
.0+1
.0+1
0.0)
IBA
+TD
Z+B
AP
(2.0
+2.0
+15.
0)
Culture room Glasshouse Wire house
Number of sprouted budsSprouting %
Fig. 5.2: Effect of different environments and media formulations on shoot forcing from
Pecan stems segments (25 cm long) during the spring season (February -
March)
75
Fig. 5.3 Fig. 5.4 Fig. 5.5 Fig. 5.6
Fig. 5.3: Pecan shoot segments immersed in forcing solution, i.e., distilled water with
200 mg/l 8-HQC, 30 g sucrose and IBA + TDZ both at a concentration of 0.5
µM and BAP (5.0 µM) under culture room conditions (1.0 x).
Fig. 5.4: An enlarged view of the marked part from Fig. 5.3 (1.0 x).
Fig. 5.5: Shoot segments immersed in distilled water with 200 mg/l 8-HQC, 30 g
sucrose, IBA + TDZ (2.0 + 2.0 µM) and BAP (15.0 µM) under culture room
conditions (1.0 x).
Fig. 5.6: An enlarged view of dotted central part from Fig. 5.5 (1.3 x).
Under wire house environment, stem segments showed a slight swelling of buds at
day 6 of initial culture (Fig. 5.7 - 5.8). The percentage sprouting observed was 37.84.
However, maximum (89.45 %) sprouting of buds was observed under glasshouse
conditions at day 6 (Fig. 5.2). Swelling of buds was more pronounced as compared to
other environments, i.e., culture room and wire house (Fig. 5.9 - 5.10). All the cultures
were clean through out the experimental period placed under glasshouse and wire house
environments.
76
Fig. 5.7 Fig. 5.8
Fig. 5.9 Fig. 5.10
Fig. 5.7: Pecan shoots immersed in glass-jars highlighting the swelling of buds (arrows)
in forcing solution containing IBA + TDZ (0.5 + 0.5 µM) and BAP (5.0 µM)
under wire house conditions at day 6 (1.0 x).
Fig. 5.8: Pecan shoots immersed in glass-jars indicating the swelling of buds appearing
green in colour (arrows) in medium containing IBA + TDZ (0.5 + 0.5 µM) and
BAP (5.0µM) in glasshouse at day 6 (1.0 x).
Fig. 5.9: Dotted area highlighting swelling of buds appearing bright green from shoots
placed in forcing solution containing IBA + TDZ (2.0 + 2.0 µM) and BAP at
15.0 µM under glasshouse conditions at day 6 (1.0 x).
Fig. 5.10: An enlarged view of highlighted area from Fig. 5.9 (2.5 x).
77
5.2 FORCING LARGE STEM SEGMENTS
Mature stem segments (diameter range; 1.0 - 4.6 cm) were cut from field grown
adult Pecan trees. Cut to yield 40 cm long logs were randomly picked for forcing
epicormic buds by placing horizontally in flats (52 × 25 × 6.5 cm; L × W × H) filled with
sterilized coccopeat, sand, sawdust and a control (empty flat). These flats were kept
under three environmental conditions, i.e., culture room, glasshouse or wire house.
A quite good response for the forcing of softwood shoots was observed from the
mature stem segments of adult field grown Pecan tree. Seasons significantly influenced
the growth of softwood shoots in different environments from logs of Pecan (Table. 5.1).
During winter season (December - January), a remarkable response in terms of sprouting
of the epicormic buds from logs was observed in all media under glasshouse conditions
(Table. 5.1). Sprouting of epicormic buds on sand, sawdust or coccopeat was initiated
after 8, 9 and 11 days (Fig. 5.11 - 5.18) respectively. The maximum number of sprouting
(16.1) with 4.6 number of shoots of 3.1 cm long having maximum number of nodes (9.1)
and leaves (8.1) was observed on sand under glasshouse conditions. The total number of
shoots per flats was also relatively greater in glasshouse with more shoot length and
number of leaves (Table. 5.1) followed by coccopeat. A most remarkable feature, i.e., the
development of inflorescence (Male flower: Catkins) was seen in logs placed on sand
under glasshouse conditions (Fig. 5.11 - 5.12). Multiple sprouting of epicormic buds was
also observed in all media under all environments (Fig. 5.13 - 5.18).
78
TABLE. 5.1: EFFECT OF DIFFERENT ENVIRONMENTS AND MEDIA TO FORCE
EPICORMIC BUDS FROM PECAN LOGS (40 CM LONG STEM SEGMENTS) DURING WINTER SEASON (DECEMBER - JANUARY)
Environmental
conditions Medium
(Sterilized)No. of
sprouts No. of shoots
Shoot length (cm)
No. of nodes
No. of leaves
Sand 5.50b 2.00c 2.1ab 4.0c 4.0c
Coccopeat 5.00b 2.0c 1.7ab 6.0b 6.0bCulture Room Sawdust 2.30d NRd NRd NRe NRe
Sand 16.10a 4.6a 3.1a 9.1a 8.1a
Coccopeat 6.23b 2.85b 2.5b 6.66b 5.5bcGlasshouse Sawdust 4.20bc 2.0bc 2.3bc 5.4bc 5.3bc
Sand 4.10cd 2.1bc 1.5d 2.0d 4.0c
Coccopeat 2.50cd 1.0cd 1.0de 1.0de 2.0dWire House Sawdust 2.30d NRd NRe NRe NRe
NR Not Recorded → The data represented the means of 9 logs per medium/ environment/ season and three logs were placed in each tray. → Different letters within a specific column represent significant difference at P< 0.05 according to Duncan’s Multiple Range Test.
79
Figure: 5.11 - 5.18: Initiation of sprouting and development of softwood shoots in
different media (coccopeat, sand or sawdust) under glasshouse conditions at 25 ±° C
Fig. 5.11 Fig. 5.12 Fig. 5.13 Fig.5.14
Fig. 5.11: Initiation of sprouting (arrows) of shoots in logs placed on sterilized sand, a
broader arrow indicating the enlarged view of the sprouted buds at day 9
(1.0 x).
Fig. 5.12: Further development of sprouted epicormic buds seen in Fig. 5.11 in to male
inflorescence, “catkin” (arrows) at day 17 (1.0 x).
Fig. 5.13: An enlarged view of log placed in sand showing sprouting of multiple buds
(1.0x).
Fig. 5.14: Multiple sprouting observed in logs placed on sterilized coccopeat, arrow
pointing towards enlarged view (1.0 x).
80
Fig. 5.15 Fig. 5.16 Fig. 5.17 Fig. 5.18
Fig. 5.15: Development of multiple shoots (arrows) in logs on sterilized coccopeat
(1.0 x).
Fig. 5.16: Emergence of multiple shoots (arrows) in logs on sterilized sawdust (1.0 x).
Fig. 5.17: Pecan logs placed in flats filled with sterilized coccopeat showing the
sprouting of buds and development of softwood shoot (highlighted
areas) (1.0 x).
Fig. 5.18: Enlarged views of the highlighted areas from Fig. 5.17.
81
During winter (December - January) season, under culture room and wire house
conditions, a fair number of epicormic buds were sprouted in all media (Table. 5.1; Fig.
5.19 - 5.21). Mostly the sprouts survived for a maximum of 6 days and afterwards
become dry. The left over survived sprouts were later on elongated into softwood shoots
in sand and coccopeat. None of the sprouted epicormic buds developed into softwood
shoots in sawdust. However, further growth and development of softwood shoots was
limited in all media.
Fig. 5.19 Fig. 5.20
Fig. 5.21
Fig. 5.19: Sprouting of epicormic buds observed in sawdust (arrows) at day 8 (1.0 x).
Fig. 5.20: Pecan logs placed in flats filled with sterilized sand showing the sprouting of
buds (arrows), at day 8 at 25 ± 2 ºC (1.0 x).
Fig. 5.21: Sprouting of epicormic buds (arrows) in coccopeat at day 8 at 25 ± 2 ºC
(1.0 x).
82
During the spring season (February - March), fair response was observed in terms
of buds sprouting (Table. 5.2). A significant number of sprouts (7.12) were observed in
sterilized sand followed by coccopeat (4.41) under glasshouse environment (Fig. 5.22).
Maximum number of shoots (3.01) with fair shoot length (2.3 cm) and number of leaves
(4.78) were also observed in sand followed by coccopeat where 2.01 mean number of
shoots had 2.3 cm mean shoot length with 1.31 mean numbers of nodes and leaves (2.61)
(Table. 5.2). Although forcing of epicormic buds was possible on all media under wire
house environment but further development was restricted during the spring season (Fig.
5.22 - 5.24).
Fig. 5.22 Fig. 5.23 Fig. 5.24
Fig. 5.22: Sprouting of epicormic buds (black arrows) and softwood shoot (brown
arrows) development from logs placed in sterilized sand at day 21(1.0 x).
Fig. 5.23: Sprouting of buds (arrows) in logs placed in sterilized sawdust at day 8
(1.0 x).
Fig. 5.24: Pecan logs placed in flats filled with sterilized coccopeat. Forcing epicormic
buds using this medium was possible as shown here at day 8 (1.0 x).
83
TABLE. 5.2: EFFECT OF DIFFERENT ENVIRONMENTS AND MEDIA TO FORCE EPICORMIC BUDS FROM PECAN LOGS (40 CM LONG STEM
SEGMENTS) DURING SPRING SEASON (FEBRUARY - MARCH)
Environmental conditions
Medium (Sterilized)
No. of sprouts
No. of shoots
Shoot length (cm)
No. of nodes
No. of leaves
Sand 4.12b 1.32c 1.5a 2.7b 4.62a
Coccopeat 3.24bc 1.15c 1.21ab 1.23bc 2.25b
Culture Room Sawdust 1.0de d c c c
Sand 7.12a 3.01a 2.3a 2.91a 4.78a
Coccopeat 4.41b 2.01b 1.9a 1.31bc 2.61bc
Glasshouse Sawdust e d c c c
Sand 2.2cd d c c c
Coccopeat 1.5cde d c c c
Wire House Sawdust 1.9cd d c
c c
→ The data represented the means of 9 logs per medium/ environment/ season and three logs were
placed in each tray. → Different letters within a specific column represent significant difference at P<0.05 according to
Duncan’s Multiple Range Test.
84
During the autumn season (August - September), forcing epicormic buds was
possible only in sterilized sand under culture room or glasshouse conditions (Table. 5.3).
Better results with regard to mean number of sprouts (5.46) and softwood shoots (3.27)
were recorded for sand in glasshouse conditions (Fig. 5.25). These shoots had 3.31 cm
length with highest mean number of nodes (5.22) and leaves (6.23). No sprouting,
however, was observed in any other medium under all the three tested environmental
conditions.
Fig. 5.25
Fig. 5.25: Sprouted epicormic buds (brown arrows) and softwood shoots (black arrows)
developed in sand at day 11 (1.0 x).
85
TABLE. 5.3: EFFECT OF DIFFERENT ENVIRONMENTS AND MEDIA TO FORCE EPICORMIC BUDS FROM PECAN LOGS (40 CM LONG MATURE STEM
SEGMENTS) DURING AUTUMN SEASON (AUGUST - SEPTEMBER)
Environmental conditions
Medium (Sterilized)
No. of sprouts
No. of shoots
Shoot length (cm)
No. of nodes
No. of leaves
Sand 4.12b 1.95b 1.72b 4.55b 4.92b
Coccopeat c c c c c
Culture Room Sawdust c c c c c
Sand 5.46a 3.27a 3.31a 5.24a 6.23a
Coccopeat c c c c c
Glasshouse Sawdust c c c c c
Sand c c c c c
Coccopeat c c c c c
Wire House Sawdust c c c c c
→ The data represented the means of 9 logs per medium/ environment/ season and three logs were placed
in each tray → Different letters within a specific column represent significant difference at P<0.05 according to
Duncan’s Multiple Range Test.
86
The shoots were green and grew more vigorously in sand than any other media
under glasshouse conditions during winter season (Fig. 5.26). Similarly the number of
shoots was also highest (4.6) from sand under glasshouse conditions. This also happened
for other environmental conditions as well. A diagrammatic representation of schematic
placement of logs and development of shoots in sand and coccopeat under glasshouse
conditions during winter season is shown below (Fig.5.26 - Fig. 5.30).
Fig. 5.26 - Fig. 5.30: Forcing epicormic buds and growth of softwood shoots
observed in Pecan logs placed in flats filled with sterilized sand, coccopeat
and sawdust under glasshouse conditions maintained at 25 ± 2 ºC during
winter season.
A C
B D
Fig. 5.26: Sprouting and growth of soft-wood shoots from logs placed in sand. A & B)
A view of flat filled with sterilized sand (1.0 x). C) An enlarged view of the
highlighted area from A (1.2 x). D) An enlarged view of the highlighted area
from B (2.1 x).
87
Fig. 5.27
Fig. 5.27: Forcing of epicormic buds and development of softwood shoots in logs
placed in flats filled with sterilized sand at day 47 (1.0 x).
Fig. 5.28 Fig. 5.29
Fig. 5.28: A softwood shoot from logs placed in sterilized sawdust at day 47 (1.0 x).
Fig. 5.29: Softwood shoots (arrows) from logs placed in flats filled with sterilized
coccopeat at day 47 (1.0 x).
88
Sawdust
Coccopeat
Sand
Fig. 5.30
Fig. 5.30: Pecan logs placed in flats filled with sterilized sand, coccopeat and sawdust
showing the sprouting and growth of softwood shoot at day 51 (1.0 x).
89
An interactive effect of season, environment or medium for forcing softwood
shoots from epicormic buds regarding the parameters studied are depicted in Table. 5.4.
The rate of epicormic bud induction (number of sprouts) was good enough (97.554) on
medium followed by season (79.570) or environment (68.883). Similarly, results were
also significant for number of shoots, shoot length, number of nodes or leaves. The
interactive effect of season and environment was significant with shoot parameters
except shoot length (0.853). The interaction of season with medium was mostly non-
significant, whereas the effective of both environment and medium was promising for
number of sprouts or shoots. It was observed that number of sprouts and leaves were
good enough as compared to other parameters as affected by the interaction of these
factors (seasons × environment × medium).
TABLE. 5.4: ANALYSIS OF VARIANCE OF DIFFERENT PARAMETERS FOR SHOOT FORCING OF PECAN SEGMENTS
Mean square
Source of variation
df Number of
Sprouts
Number of
Shoots
Shoot Length
(cm)
Number of
Nodes
Number of
Leaves Season (A) 2 79.570* 11.297* 7.667* 66.216* 48.465* Environment (B) 2 68.883* 11.237* 11.624* 60.758* 51.548* Medium (C) 2 97.554* 15.252* 9.013* 29.865* 52.465* A × B 2 18.835* 3.144* 0.853NS 28.222* 9.068* A × C 2 12.098* 1.085NS 8.835NS 4.497NS 1.407NS
B × C 4 32.053* 2.680* 0.547NS 5.465NS 5.594NS
A × B × C 4 7.071* 0.747NS 1.117NS 3.392NS 7.729* Error 60 2.193 0.883 0.897 2.420 2.534
* indicates significant or NS non-significant at P<0.05 value according to F-test.
90
An overall effect of seasons (Autumn, Spring and Winter) on epicormic bud
induction potential with reference to bud-derived shoot parameters is shown in Fig. 5.31.
The total number of sprouts (6.89) per flat was greater during winter followed by spring
season (5.24). Although, the number of sprouts was higher but few (3.9) were elongated
to produce softwood shoots, during winter season. Maximum number of nodes (6.07)
and leaves (5.49) were observed with maximum shoot length of 3.21 cm during the
winter season (Fig. 5.31).
efdefdefdef
cde
defcde
bcbcabc
cd bcd
abcab
a
0
1
2
3
4
5
6
7
8
No. of sprouts No.of shoots Shoot length No. of nodes No. of leaves
Growth parameters
Num
ber/l
engt
h of
sho
ot
Autumn Spring Winter
Fig. 5.31: Effect of different seasons on epicormic bud induction potential with reference
to bud-derived shoot parameters in Pecan logs. Vertical bars above the
columns are the SE (±) of the means. Different letters above the vertical bars
representing the significant differences according to Duncan’s Multiple Range
test at P<0.05 value.
91
An overall effect of media (Coccopeat, Sand and Sawdust) on epicormic bud
induction potential with specific reference to bud-derived shoot parameters is shown in
Fig. 5.32. Amongst all three tested media, sand was quite promising with higher mean
number of sprouts, shoots, leaves or shoot length. The mean number of sprouts was
highest (8.49) followed by the number of nodes (6.1) or leaves (5.33) per flat tray on
sand as compared to other tested media. However, the number of shoots was low (4.15)
with 3.9cm mean shoot length (Fig. 5.32).
def def
bcd bcd
bc bcbc
abab
a
defdefbcd
bcbc
0
1
2
3
4
5
6
7
8
9
10
No. of sprouts No.of shoots Shoot length No. of nodes No. of leaves
Growth parameters
Num
ber/L
engt
h of
sho
ot
Coccopeat Sand Saw dust
Fig. 5.32: Effect of different media on epicormic bud induction potential with reference
to bud-derived shoot parameters in Pecan logs. Vertical bars above the
columns are the SE (±) of the means. Different letters above the vertical bars
representing the significant differences according to Duncan’s Multiple Range
test at P<0.05 value.
92
The three different environmental conditions also greatly affected the forcing
potential of the logs. An overall effect of environment (Culture room, Glasshouse and
Wire house) with respect to epicormic bud induction potential is shown in Fig. 5.33. The
total number of sprouts (6.92) per flat was greater in glasshouse conditions followed by
culture room (3.18) and wire house (2.41). Similarly, more numbers of shoots (4.005)
were developed in glasshouse than culture room conditions. Although the maximum
shoot length obtained was 2.56 cm but number of nodes (5.28) and leaves (5.26) were
quite high in glasshouse. However, results were not satisfactory under wire house
conditions (Fig.5.33).
cdefcdef
bcdbcd
bc
cde
bcd
abab
a
fefefef
cde
0
1
2
3
4
5
6
7
8
No. of sprouts No.of shoots Shoot length No. of nodes No. of leaves
Growth parameters
Num
ber/L
engt
h of
sho
ots
Culture room Glasshouse Wire house
Fig. 5.33: Effect of different environments on epicormic bud induction potential
with reference to bud-derived shoot parameters in Pecan logs. Vertical
bars above the columns are the SE (±) of the means. Different letters
above the vertical bars representing the significant differences
according to Duncan’s Multiple Range test at P<0.05 value.
93
The cumulative forcing potential of the logs regarding the parameters studied
(number of sprouts, shoots, nodes, leaves and shoot length) under the influence of media,
environment, and season has been shown in Fig. 5.34. It is clear that greater number of
epicormic buds (6.425) were forced but lesser numbers of them were elongated to
produce softwood shoots (3.146) with mean shoot length of 2.978 cm. These softwood
shoots have 6.838 numbers of nodes followed by 5.830 numbers of leaves during the
present study.
c
c
ab
aa
0
1
2
3
4
5
6
7
8
No. of sprouts No.of shoots Shoot length No. of nodes No. of leaves
Growth parameters
Num
ber/L
engt
h of
sho
ots
Fig. 5.34: A cumulative effect of media, environment and season on forcing potential of
the logs regarding the parameters studied (number of sprouts, shoots, nodes,
leaves and shoot length) in Pecan. Vertical bars above the columns are the SE
(±) of the means. Different letters above the vertical bars representing the
significant differences among different values according to Duncan’s Multiple
Range test at P<0.05 value. This figure depicts the cumulative data of three
experiments. In each experiment 9 logs were placed in three media (each
medium has 3 trays and 3 logs per tray) under three environmental conditions
during three seasons.
94
A striking feature related to log diameter was also observed during forcing of
large stem segments. A variation in forcing response was observed in relation to log
diameter (1.0 - 5.0 cm). Maximum forcing of epicormic buds was recorded in logs with
diameter < 2.5 cm under all environmental conditions, whereas logs with diameter ≥ 2.5
cm showed better response in terms of development and growth of softwood shoots.
A noteworthy phenomenon, i.e., the formation of callus was also observed at the
cut surfaces from the logs of larger diameter (> 2.5 cm) during the winter season. Callus
was developed in sand and coccopeat medium under glasshouse conditions (Fig. 5.35 -
5.39). Callus formation was also observed in sand under culture room conditions (Fig.
5.40 - 5.41). No callus formation was observed under wire house environment in any
medium used.
Fig. 5.35 Fig. 5.36 Fig. 5.37
Fig. 5.35: Pecan logs placed in sand indicating the formation of callus (dotted area) at
the cut surfaces under glasshouse conditions at day 17 (1.0 x).
Fig. 5.36: An enlarged view of the dotted portion from Fig. 5.35 (1.6 x).
Fig. 5.37: Another photograph showing the formation of callus (arrows) at the cut
surface of log placed in sand under glasshouse conditions (1.0 x).
95
Fig. 5.38 Fig. 5.39 Fig. 5.40 Fig. 5.41
Fig. 5.38: Pecan log indicating the formation of callus (arrows) at the cutting points in
coccopeat under glasshouse conditions.
Fig. 5.39: An enlarged view of the highlighted portion from Fig. 5.38 (1.0 x).
Fig. 5.40: Pecan logs placed in sand indicating the formation of callus (an arc) at the cut
surfaces in sand under culture room conditions (1.0 x).
Fig. 5.41: An enlarged view of the highlighted portion from Fig. 5.40 (1.0 x).
96
A major problem associated with Pecan tissue culture is the culture
contamination. Preventing or avoiding contamination of plant tissue culture is critical to
successful micropropagation. For this reason a number of other methods of
micropropagation have been developed for recalcitrant tree crops. These methods utilize
different media, i.e., perlite, vermiculite, peat moss etc. In the present research work,
coccopeat, sand or sawdust was used. However, fungal contamination was also observed
in the media (Coccopeat and Sawdust) after two weeks of initial cultures under culture
room conditions. This type of contamination was found developing as a whitish fungal
mat all over the medium surface within the vicinity of logs (Fig. 5.42). It also extends
between the logs towards the cut surfaces (Fig. 5.43) in coccopeat. Fungal contamination
also affected the growing points of logs, i.e., the sprouting epicormic buds (Fig. 5.44).
On the other hand, it was found as whitish globular structures over the surface of sand
(Fig. 5.45). To overcome the problem of culture contamination, 0.18 percent hydrogen
peroxide (H2O2) solution was sprayed manually once a day.
Fig. 5.42 Fig. 5.43
Fig. 5.42 - 5.43: Pecan logs placed in sawdust, dotted areas indicating the presence of
contamination on the media surfaces under culture room conditions (1.0 x).
97
Fig. 5.44 Fig. 5.45
Fig. 5.44: Pecan logs placed in sawdust, arrows indicating the contamination of sprouted
buds under glasshouse conditions (1.0 x).
Fig. 5.45: Pecan logs placed in sterilized sand, the highlighted dotted areas indicating the
presence of contamination on the media surfaces under culture room
conditions (1.0 x).
98
5.2.1 ESTABLISHMENT OF SOFTWOOD SHOOTS IN DIFFERENT
ROOTING MEDIA
After acquiring sufficient length (≥ 4 cm long), the softwood shoots were cut with
the help of sharp razor and kept in a beaker containing water to prevent dehydration and
were surface disinfested by rinsing 5 times with autoclaved distilled water. These
softwood shoots were then treated with 1000 or 2000 ppm NAA or IBA or a combination
of IBA + NAA (1000 + 1000 ppm) by dipping in PGRs solution for 10 seconds.
Afterwards, with a sterilized forcep these pre-treated softwood shoots were planted in
flats (52 × 25 × 6.5 cm; L × W × H) filled with sterilized sand (Fig. 5.46 - 5.47). Pre-
treated softwood shoots were also transferred for rooting to pots (9.5 × 12 cm) containing
peat moss and vermiculite (Fig. 5.48 - 5.53). All pots and flats were irrigated very
carefully with squeezing bottle and kept in culture room at 25 ± 2ºC temperature in 16 h
photoperiod for further growth. The softwood shoots did not show any rooting, got
contaminated and ultimately became necrotic (Fig. 5.54 - 5.56). Contamination of
softwood shoots was observed in peat moss and vermiculite medium (Fig. 5.54 - 5.55).
On the other hand, no signs of contamination were observed in sand, but rooting was not
initiated.
Fig. 5.46 Fig. 5.47
Fig. 5.46 - 5.47: Soft wood shoots derived from epicormic/ latent buds placed in
sterilized sand for rooting phenomenon under culture room environment
at 25 ± 2 ºC (1.0 x).
99
Fig. 5.48 Fig. 5.49 Fig. 5.50
Fig. 5.48: A soft wood shoot harvested from forced Pecan logs placed in peat moss for
rooting under controlled environmental conditions (1.0 x).
Fig. 5.49 - 50: Soft wood shoots harvested from forced Pecan logs placed in different
grades of vermiculite for rooting under controlled environmental
conditions (1.0 x).
Fig. 5.51 Fig. 5.52
Fig. 5.51: A soft wood shoot harvested from forced logs with the leaves removed placed
in vermiculite for rooting under controlled environmental conditions (1.0 x).
Fig. 5.52: A comparison of different rooting media with softwood shoots maintained in
culture room at 25 ± 2 ºC (1.0 x).
100
Fig. 5.53 Fig. 5.54 Fig. 5.55
Fig. 5.56
Fig. 5.53: Plastic pots containing softwood shoots were placed under an artificially
constructed chamber with transparent polyethylene sheet for the maintenance
of high humidity kept in culture room at 25 ± 2 ºC (1.0 x).
Fig. 5.54: A plastic pot showing fungal contamination (arrow) along the base of dried
softwood shoot kept in culture room at 25 ± 2 ºC (1.0 x).
Fig. 5.55: A plastic pot showing fungal contamination spread on medium surface along-
with dried shoot in the center kept in culture room at 25 ± 2 ºC (1.0 x).
Fig. 5.56: Dried softwood shoot showing no development of roots with yellowish fungal
contamination at base (1.0 x).
101
DISCUSSION
A number of important ornamental tree species can be propagated by softwood
stem cuttings (Henry and Preece, 1997a, b). This technique was also employed for
recalcitrant woody species (Preece and Read, 2003; Aftab and Preece, 2007; Akram and
Aftab, 2009). Softwood shoots can be forced from woody stems through two basic
means (Preece and Read, 2007). For example, Read and Yang (1991) used a forcing
solution to stimulate tip growth from dormant woody stems and subsequently used
shoots both for tissue culture and soft wood cuttings. Alternatively, larger stem segments
can be cut and placed in a suitable greenhouse medium and epicormic/ latent buds can be
forced to grow (Preece and Read, 2007). Forcing large stem segments of woody plants is
a way to stimulate epicormic (dormant, latent or suppressed) buds to grow into softwood
shoots (Henry and Preece, 1997a, b; Preece et al., 2002; Preece and Read, 2003, 2007;
Preece, 2008). Usually, large branches are cut from the lower portion (more juvenile) of
the woody plants (Preece and Read, 2003). The resulting forced softwood shoots can be
used as stem cuttings for rooting (Henry and Preece, 1997b) or as an explant sources for
micropropagation (Preece, 2003; Mansouri and Preece, 2009). Shoot forcing as well as
forcing epicormic buds is well documented for several temperate plants (Van Sambeek et
al., 2002; Preece and Read, 2003; Aftab et al., 2005). The potential of these forcing
methods for micropropagation of Pecan has never been investigated in detail previously
though, Aftab and Preece (2007) reported a preliminary work on forcing epicormic/
latent buds from large stem segments of Pecan. In their work, a mean number of 3
harvestable shoots per log was obtained. A very high microbial contamination limited
their success under fog or mist conditions. Nevertheless, they demonstrated that shoot
forcing offers a strong possibility to raise tissue to be utilized under in vitro conditions.
The results of the present research work have shown faster bud break from the terminal
102
shoots cut during the dormant season and placed in forcing solution under glasshouse
conditions. Forcing solution was used to extend the season by forcing woody stems
during the dormant period (Read et al., 1984; Yang et al., 1989; Read and Yang, 1989).
Furthermore, Read and Yang (1987) demonstrated forcing solution to be a means of
providing PGRs to the forced tissues. Read and Yang (1988) in a study involving liliac
and privet, reported that bleach treatment increased percent bud break and shoot length
while reducing days to bud break. Yang and Read (1992) also reported faster bud break
and more bud and shoot elongation if the cut stems were treated with bleach solution for
15 minutes prior to forcing. During the present investigation, although faster bud break
was observed but afterwards such buds turned brown and became harder at day 13.
Subsequent development into softwood shoots was also not achieved.
During the present research work, three media (coccopeat, sand and sawdust)
were tested for forcing large stem segments of Pecan. Previously, the use of perlite and
vermiculite as a greenhouse medium was reported for shoot forcing of large stem
segments in temperate woody dicots (Preece and Read, 2003; Aftab et al., 2005; Aftab
and Preece, 2007). Forcing medium in this study significantly influenced the production
of softwood shoots from large stem segments of Pecan. Amongst the three media tested,
sterilized sand had a pronounced effect on epicormic bud induction/sprouting and
subsequent elongation of softwood shoots. Maximum softwood shoots (2.92) were
observed in sterilized sand followed by coccopeat (1.80) or sawdust (1.6). Generally,
Pecan grows best on well-drained sandy loam or loamy sand that are not subject to
prolonged flooding (Andersen and Crocker, 2009). Sand is a naturally occurring granular
material consisting of silica (Silicon dioxide, or SiO2), which, because of its chemical
inertness is preferred as growth medium. Sand is highly porous having excellent drainage
characteristics thus providing most suitable environment for the shoot forcing in Pecan.
103
In addition, sand can not absorb or adsorb any organic substances, toxic or inhibitory
secondary metabolic products are washed from plantlets, if the flask contents are shaken
briefly under in vitro cultures. Coccopeat stands second for the production of softwood
shoot from large stem segments of Pecan. Coccopeat, also known as coir pith, coir dust,
or simply coir, is a natural, and renewable resource produced from coconut husks by the
coconut industries. It consists of coarse fiber, lignin and cellulose material. It has high
water holding capacity thus used as potting medium for plant propagation in many
countries (http://www.en.wikipedia.org/wiki/Coconut). However, it has only been
reported as rooting substrate to propagate temperate and tropical species. Its high water
retention capacity might be the reason for lower number of production of softwood
shoots in Pecan. On the other hand, sawdust was found to be poorest in terms of forcing
of epicormic/ latent buds from large stem segments. During the present investigation,
sawdust was locally obtained form the timber market of Lahore. It was composed of fine
particles of different woods, such as Cedrus deodara, Dalbergia sisso, Acacia nilotica,
Salmalia malabarica or Eucalyptu spp. The exact composition of the sawdust was
unknown, however, Cedrus, Dalbergia, Acacia, Eucalyptus or Salmalia were found in
different ratios in different samples obtained from time to time. Several organic
compounds are naturally found in woody tree species. These organic compounds are
generally termed as secondary metabolic products such as, alkaloids, terpenoids, tannins,
phenolics and several essential oils. It was previously reported that both physical and
chemical properties of the medium play a vital role for normal plant growth and
development (Ahmed et al., 1996). In sawdust medium, the presence of such toxic
compounds may restrict the development of epicormic buds to grow out as softwood
shoots from large stem segments of Pecan. The results of present investigation have
shown that use of a mixture of different woods as a medium supplement was poor in
104
terms of forcing epicormic buds in Pecan. This suggests that sawdust derived from an
individual tree should also be tested to check the effect on growth and production of
softwood shoots from Pecan logs. Previously, the use of sand, coccopeat or sawdust was
reported as propagation media in greenhouse or nursery (Abu-Rezq, 2009; Mhango et
al., 2008; Gungula and Tame, 2007; Bugbee, 1999). Several authors mainly from
developing countries (Babbar and Jain, 1998; Naik and Sarkar, 2001; Mohan et al.,
2004) have been looking forward for a variety of low-cost agar substitute for
micropropagation studies. Recently, silica sand was used as medium supplement in the
micropropagation studies of woods (Prknova, 2007). Nonetheless, their use as a medium
for forcing epicormic buds in woody plants has never been investigated prior to this
study. The use of sand, coccopeat or sawdust as a propagation media was preferred in
order to cut down expenses in using imported relatively expensive growing media
(perlite or vermiculite). Furthermore, this technique is also cheap since the materials
needed for the media could be sourced locally and readily available.
During the present investigation, three environmental conditions (culture room,
glasshouse or wire house) were also compared demonstrating a significant role for
forcing softwood shoots from epicormic buds of large stem segments of Pecan. Highest
(2.92) mean number of softwood shoots were obtained under glasshouse conditions
without mist or fog system. Although maximum sprouting of epicormic buds was
achieved, yet, there was a high percentage of visible buds that failed to elongate
suggesting that there was indeed a potential for the production of many more harvestable
shoots. Most of the visible buds did not elongate sufficiently to make harvestable shoots,
instead resulted in short sprouts (≤ 1.5 cm) that could be utilized as explants for
micropropagation. During the present investigation, mean number of softwood shoots
(2.92) seems to be relatively lower than other tree species of temperate origin (Preece
105
and Read, 2003; Aftab et al., 2005; Aftab and Preece, 2007; Preece and Read, 2007).
Henry and Preece (1997a) reported a higher (6.9) number of softwood shoots from Acer
species (Red maple). Similarly, in another study involving silver maple, Aftab et al.,
(2005) reported 6.7 mean number of softwood shoots under mist conditions. The
production of high number of softwood shoots in previous reports might be due to the
established ideal forcing conditions. It was previously reported that intermittent mist was
a more effective forcing environment for epicormic softwood shoot forcing (Preece et
al., 2003; Preece and Read, 2003). However, high microbial contamination was observed
with the explants derived from softwood shoots forced under mist or fog (Preece and
Read, 2003). The results of the present study were quite promising in terms of
production of relatively fair number of softwood shoots under glasshouse conditions
without mist or fog. Thus, the present study demonstrates an efficient and quite cheaper
method for forcing softwood shoots in Pecan under glasshouse conditions with reduced
rate of microbial contamination.
In addition, seasons also influenced the production of softwood shoots from large
stem segments (Preece and Read, 2003). The results from the present investigation have
shown that bud break was not possible at all during July to September (summer), but was
rapid enough during December - January (winter). Highest mean numbers of softwood
shoots were produced during the winter as compared to spring or autumn seasons. In
contrast, Van Sambeek et al., (1997a) reported that branch segments from eastern black
walnut (another member of the family, Juglandaceae) collected in March from Illinois,
USA, produced softwood shoots after receiving the chilling treatment during the
preceding September. This could happen because September is the dormant season for
eastern black walnut and dormancy had been initiated on the latent buds. However
during the present investigation, highest mean numbers of softwood shoots were
106
produced during December (begining of winter). It might be due to the fact that the
prevailing high relative humidity and low temperature during the winter season have
played a significant role in the forcing of epicormic buds. Maximum production of
softwood shoots during winter season may be attributed to the fact that Pecan grows well
in temperate environments. Usually, the optimum temperature for various growth
parameters of temperate tree species is lower (Pijut and Moore, 2002) than the tropical
tree species (Aftab and Preece, 2007). In addition, seasonal variations in shoot forcing of
temperate woody plants are well documented (Preece, 2003) and seems to hold for Pecan
as well.
The effect of stem diameter on shoot forcing potential was also determined.
Maximum epicormic bud induction was observed in logs with diameter < 2.5 cm but
further elongation into harvestable softwood shoots was restricted. Usually, the
epicormic bud induction was good in logs with diameter ≥ 2.5 cm and the developing
softwood shoots grew more vigorously. Length of softwood shoots was also greater on
the large diameter stem sections. This might be due to the presence of higher amount of
reserved food material (Preece and Read, 2003). Larger diameter stem segments contain
higher concentration of carbohydrates and other growth substances that are usually
necessary for softwood shoot production (Henry and Preece, 1997b). Stem segments
with smaller diameters perhaps have less stored food which results in poorest shoot
forcing (Preece and Read, 2007). Furthermore, it was previously demonstrated that stem
diameter was less important than stem length (Henry and Preece, 1997b) because larger
stem segments had more stored food than shoot tips that may contribute to enhanced
softwood shoot growth (Preece and Read, 2003). The diameter of the logs has also been
related with the age of the source plant (Henry and Preece, 1999). The large stem
segments offer an advantage over the smaller ones as they can be forced for an extended
107
period of time of the year. Generally, stem segments should be of at least 2.5 cm
diameter (Preece and Read, 2007). During the present investigation softwood shoot
forcing was also observed from stem segments having diameter even greater than 5.6 cm.
Logs with small diameter range (< 2.5 cm) were less responsive. The upper log diameter
limit for forcing stem segments is unknown and it seems to be species-specific (Henry
and Preece, 1997b) and possibly tree age specific (Van Sambeek et al., 2002).
Nonetheless, Fishel et al., (2003) suggested that there may be no upper limit to stem
diameter for forcing because the lowest segments of main stems produced the most
shoots.
Rooting of woody species is usually more difficult than that of herbaceous plants.
Kyte and Kleyn (1999) emphasized that woody species are more difficult than herbs in
cultures in vitro, and conifers are especially more difficult than deciduous woody
species. Some species form roots easily, others are recalcitrant, such as Pecan. During
the present investigation, the availability of less number of softwood shoots with
considerable shoot length was the limitation that barred from applying the experiments
under in vitro conditions on large scale. Therefore, Pecan softwood shoots (≥ 4 cm) were
subjected to different greenhouse rooting media (sand, peat moss or vermiculite), but no
significant rooting was recorded. Fett-Netto et al., (2001), suggested that woody species
lose their rooting capacity with seedling age. Difficulties in root induction are also
related to juvenility or maturity of the shoots (Ballester et al., 1999).
During the present investigation, culture contamination was a major problem that
limited the production of softwood shoots on large scale. Contamination was observed in
media (sand or coccopeat) under glasshouse and culture room conditions due to a high
relative humidity and temperature. Trichoderma is a naturally occurring fungus in
coccopeat, however, it is not present in sterilized coccopeat. Under humid conditions the
108
medium invariably got contaminated. Contamination was reduced to 25 % by the
application of 0.18 % hydrogen peroxide solution (Aftab et al., 2005) on a daily basis.
Furthermore, establishment of Pecan softwood shoots under in vitro or ex vitro
conditions was also quite a challenge because of culture contamination. Softwood shoots
forced under natural or glasshouse conditions are susceptible to microbial attack on the
surfaces of softwood shoots (Preece and Read, 2003). Under humid conditions the
medium got contaminated (30 - 40 %). A 100 % contamination rate was a major
limitation during rooting of softwood shoots. Without the use of fungicide, a high
percentage of the softwood cuttings were quickly infected with fungi and black shoot tip
necrosis. The softwood shoots were immersed in a solution of 0.3 % fungicide (Dithane,
M- 45) prior to transfer to rooting medium. Afterwards the fungicide solution was also
sprayed twice a day, however, the results were not satisfactory. Although our rooting
results have been discouraging in Pecan, successive rooting of cuttings from greenhouse-
forced epicormic sprouts have been reported for several other woody species of maple
(Acer saccharinum, Acer rubrum, Acer palmatum etc.), white ash (Fraxinus americana)
and European birch, Betula pendula (Cameron and Sani, 1994; Henry and Preece, 1997a;
Van Sambeek et al., 1998b).
During the present study, callus formation was observed at the cut ends of the
logs during the winter season. It was seen in sand under glasshouse and in sand or
coccopeat medium under culture room conditions. Tree responses to injuries include
formation of callus tissue (Schweingruber, 2001) at wounded section. Usually, in trees,
callus was formed in response to several injuries or wounds due to many causes
including, broken branches, abrasions and scrapes, animal damage, insect attack or fire
etc. (http://www.utextension.utk.edu/publications/spfiles/SP683.pdf). Trees strive to
isolate the damaged tissue from the outside by forming callus tissues by activating their
109
internal defense mechanism. During the present study, logs were cut and inner tissues
were exposed to the outside so the active meristematic cells proliferated and resulted in
the formation of callus at the cut, injured portion of the logs.
In conclusion, the present investigation demonstrated that forcing large stem
segments of Pecan holds promise for the production of softwood shoots under glasshouse
conditions. This simpler and cost effective method also offers a longer growing period
through the use of dormant season to produce an ample quantity of plant material
throughout the year in enhancing the micropropagation and clonal multiplication of
Pecan. During the present study, use of silica sand for softwood shoot production proved
to be quite satisfactory amongst the tested media. It came up to be a relatively much
cheaper alternative forcing medium that sustained a reasonable softwood shoot
production from large stem segments of Pecan. Further testing of silica sand as a forcing
medium is therefore recommended for other temperate or tropical woody tree species as
well.
110
CHAPTER 6
CALLUS INDUCTION AND ORGANOGENESIS
RESULTS
The objective of the present part of this research work was to sort out the best
medium as well as explant source for in vitro callus induction in Pecan (Carya illinoensis
(Wangenh.) C. Koch). It further aimed to establish a method for subsequent callus
maintenance and plant regeneration in Pecan. For this purpose, three different agar-
solidified basal medium formulations were used, i.e., DKW (Driver and Kuniyuki,
1984), MS (Murashige and Skoog, 1962) or WPM (Woody Plant Medium of McCown
and Lloyd, 1981) containing various levels and combinations of Naphthalene Acetic
Acid (NAA), 2, 4-dichlorophenoxyacetic acid (2, 4-D), or Thidiazuron (N-phenyl-N'-1,
2, 3-thidiazol-5-yl-urea; TDZ). Different explants used during this study included
immature fruit (cotyledonary portions), and mature (brown) bark (0.5 - 1.0 cm) of Pecan.
6.1 CALLUS INDUCTION FROM BARK SEGMENTS
Mature bark explants did not show any response in terms of callus induction in
MS medium supplemented with 2, 4-D (4.52 µM) or TDZ (1.0 µM) and a combination
of both 2, 4-D and TDZ, 1.0 + 1.0 µM (Table. 6.1; Fig. 6.1 - 6.2). However, MS medium
containing 13.57 or 22.61 µM 2, 4-D resulted in the induction of translucent, watery and
compact callus after which no further growth response was observed (Fig. 6.3 - 6.4). On
MS medium supplemented with TDZ (50 or 100 µM), the upper surface of bark
segments became rough in texture, cracked in center and greenish-white, compact and
111
creamy-white, translucent, watery callus was initiated from the ruptured portions of the
bark (Fig. 6.5 - 6.6) after 32 and 33 days of initial culture respectively (Table. 6.1).
Figure 6.1 - 6.6: Callus induction from bark explants cultured in MS medium placed under 16 h photoperiod at 25 ± 2 °C
Fig. 6.1 Fig. 6.2 Fig. 6.3
Fig. 6.1: Mature bark cultured on MS medium supplemented with 4.52 µM 2, 4-D
(2.5 x).
Fig. 6.2: An enlarged view of Fig. 6.1 (3.1 x).
Fig. 6.3: Bark explants cultured on MS medium supplemented with 13.57 µM 2, 4-D
showing the induction of watery, translucent callus (arrows) at day 30 of initial
culture (3.1 x).
112
cp
Fig. 6.4 Fig. 6.5 Fig. 6.6
Fig. 6.4: Mature bark cultured on MS medium supplemented with 22.61 µM 2, 4-D
showing the induction of watery, translucent callus (arrow) at day 26 of initial
culture (3.1 x).
Fig. 6.5: Induction of greenish-white, compact callus (arrows) from the cracked portions
(cp) of the mature bark on MS medium supplemented with 50 µM TDZ, at day
32 of initial culture (3.1 x).
Fig. 6.6: Creamy-white, watery, translucent, compact callus (arrows) from the ruptured
portions of the bark on MS medium supplemented with100 µM TDZ, at day 33
of initial culture (4.0 x).
113
TABLE: 6.1 EFFECT OF DKW, MS AND WPM MEDIUM WITH DIFFERENT LEVELS OF 2, 4-D, TDZ AND NAA ON CALLUS INDUCTION FROM MATURE BARK EXPLANTS OF
PECAN
Medium
Growth regulators
Concentrations (µM)
Callus induction (%)
Morphological features
4.52 67.00 (25) ± 2.51b Olive-green, compact 13.57 54.66 (27) ± 2.60c Greenish-yellow, compact
2, 4-D
22.61 49.66 (27) ± 1.45c Olive-green, compact 1.0 89.88 (26) ± 1.83a Whitish-brown, friable,
translucent, granular 50.0 90.18 (24) ± 1.28a Whitish-brown, friable, granular,
translucent
TDZ
100 89.25 (24) ± 0.74a Greenish-yellow, friable, granular
DKW
2, 4-D + TDZ
1.0 +1.0
93.70 (26) ± 3.53a Whitish-brown, watery friable, granular, embryogenic
4.52 NR*g NR 13.57 11.26 (30) ± 3.58f Translucent, watery, compact
2, 4-D
22.61 36.61 (26) ± 6.56d Translucent, watery, compact 1.0 NRg NR 50.0 24.64 (32) ± 3.41e Greenish-white, compact
TDZ
100 21.85 (33) ± 3.24e Creamy-white, translucent, watery
MS
2, 4-D + TDZ 1.0 + 1.0 NRg NR 4.52 NRg NR 13.57 2.82 (30) ± 0.70g Greenish-yellow, compact
2, 4-D
22.61 NRg NR 1.0 49.66 (25) ± 2.90c Greenish-yellow, compact 50.0 65.33 (24) ± 6.00b Transparent, compact, nodular
TDZ
100 64.33 (24) ± 4.37b Greenish-yellow, compact, nodular
WPM
2, 4-D + TDZ 1.0 + 1.0 85.77 (37) ± 0.22a Yellowish-green at top, brown at
base, friable, granular, embryogenic
NR*: Not Recorded
→Each value represents mean (± standard error) of three callus cultures and the experiment was repeated thrice. →The values in parenthesis in column for callus induction (%) are the number of days to callus induction. →Values with different letters within a specific column represent significant difference at P<0.05 according to Duncan’s Multiple Range Test.
114
Amongst WPM, the one supplemented with 2, 4-D + TDZ (1.0 + 1.0 µM)
resulted in maximum (85.77 %) callus induction after 37 days. The callus was yellowish-
green at top and brown at the base. It was granular, friable in texture and embryogenic in
nature (Fig. 6.7). As with MS medium, in WPM supplemented with different levels of
TDZ (1.0, 50 or 100 µM) the upper surface of bark segments likewise became rough in
texture, somewhat cracked and resulted in the induction of callus through such splitted
portions of bark. Compact, transparent and nodular callus (Fig. 6.8) was observed on
WPM (50 µM TDZ) and greenish-yellow, compact nodular callus was initiated on WPM
supplemented with 100 µM TDZ from the split portions of the bark (Fig. 6.9 - 6.10) after
24 days of initial culture. WPM supplemented with 13.57 µM 2, 4-D showed induction
of greenish-yellow compact callus (Fig. 6.11) at day 30 of initial culture after which
callus cultures became brown at the upper surface (Fig. 6.12). Although callus induction
was observed on WPM, further proliferation of callus cultures on all the tested levels of
2, 4-D or TDZ was limited.
Fig. 6.7 Fig. 6.8 Fig. 6.9
Fig. 6.7: Yellowish-green, granular, friable and embryogenic callus on WPM containing
2, 4-D + TDZ (1.0 + 1.0 µM) at day 37 of initial culture (1.3 x).
Fig. 6.8: Formation of compact, transparent and nodular callus (arrows) from the
cracked portions of the mature bark cultured on WPM containing 50 µM TDZ
at day 24 of initial culture (4.0 x).
Fig. 6.9: Mature bark cultured on WPM containing 100 µM TDZ showing the formation
of greenish-yellow, compact nodular callus at day 24 of initial culture (3.1 x).
115
Fig. 6.10 Fig. 6.11 Fig. 6.12
Fig. 6.10: Greenish-yellow, compact callus (arrows) induced on WPM containing1.0
µM TDZ at day 33 of initial culture (3.1 x).
Fig. 6.11: Greenish-white, compact callus formed on WPM containing 13.57 µM 2, 4-D
at day 30 of initial culture (1.7 x).
Fig. 6.12: Greenish-white, compact callus (arrow) formed on WPM containing 13.57
µM 2, 4-D at day 57 of initial culture (2.1 x).
116
After an average of 27 days of culture, callus initiation was observed from the
immature bark explants (Table. 6.1) on all the tested DKW media containing TDZ and 2,
4-D. Maximum (93.70 %) callus induction and proliferation with whitish-brown,
granular and friable morphology (Table. 6.1; Fig. 6.13 - 6.14) was observed on DKW
medium supplemented with 2, 4-D + TDZ (1.0 + 1.0 µM) followed by 90.18 % in DKW
medium containing 50 µM TDZ (Fig. 6.15). DKW medium supplemented with 2, 4-D
(4.52, 22.61 µM) resulted in olive-green, compact callus while with 13.57 µM 2, 4-D
greenish-yellow, compact callus was observed (Fig. 6.16 - 6.21). Greenish-yellow,
compact callus was observed on DKW medium containing 100 µM TDZ (Fig. 6.22) as
compared to whitish-brown, friable callus on DKW medium supplemented with 1.0 or
50 µM TDZ, respectively (Fig. 6.23 or Fig. 6.15)
Fig. 6.13 Fig. 6.14 Fig. 6.15
Fig. 6.13: Whitish-brown, granular and friable callus on DKW medium
supplemented with 2, 4-D + TDZ (1.0 + 1.0 µM) at day 36 of initial
culture (3.1 x).
Fig. 6.14: Whitish-brown, granular and friable callus on DKW medium
containing 2, 4-D + TDZ (1.0 + 1.0 µM) at day 36 of initial culture (3.1 x)
Fig. 6.15: Whitish-brown, friable callus on DKW medium supplemented with 50
µM TDZ at day 39 of initial culture (3.1 x).
117
Fig. 6.16 Fig. 6.17 Fig. 6.18 Fig. 6.19
Fig. 6.16: Olive-green, compact and nodular callus on DKW medium containing
4.52 µM 2, 4-D at day 39 of initial culture (2.9 x).
Fig. 6.17-18: Olive-green, compact, nodular callus with white luster on DKW
medium containing 22.61 µM 2, 4-D at day 39 of initial culture (2.9 x).
Fig. 6.19: Greenish-yellow compact callus on DKW medium supplemented with
13.57 µM 2, 4-D (3.1 x)
Fig. 6.20 Fig. 6.21 Fig. 6.22 Fig. 6.23
Fig. 6.20: Greenish-yellow, compact and granular callus indicating bark
remnants (arrow) on DKW medium containing 13.57 µM 2, 4-D at day 39
of initial culture (2.9 x).
Fig. 6.21: Greenish-yellow, compact callus on DKW medium containing 13.57
µM 2, 4-D at day 51 of initial culture (3.1 x).
Fig. 6.22: Greenish-yellow, friable callus on DKW medium containing 100 µM
TDZ at day 39 of initial culture (3.1 x).
Fig. 6.23: Whitish-brown, friable callus on DKW medium supplemented with 1.0
µM TDZ at day 57 of initial culture (3.1 x).
118
Maximum growth and proliferation of callus cultures was observed in all the
tested DKW media. Calluses obtained from all DKW media were maintained up to 4th
subculture by transferring them onto the respective fresh medium after every 15 day.
Further maintenance of such callus cultures was not possible due to hard-to-control
browning and necrosis that resulted in sudden callus death (Fig. 6.24 - 6.26). During
each subculture, these calluses were also transferred on MS medium supplemented with
BAP (2.22 µM) or BAP + TDZ (2.22 + 1.0 µM) for plant regeneration. Plant
regeneration, however, was not possible during the present investigation.
Fig. 6.24 Fig. 6.25 Fig. 6.26
Fig. 6.24: Initiation of browning of greenish-yellow, granular compact callus on
DKW medium supplemented with 100 µM TDZ at day 57 of initial
culture (3.1 x).
Fig. 6.25: Browning of callus after 4th subculture on DKW medium containing
13.57 µM 2, 4-D (3.1 x).
Fig. 6.26: Browning of callus after 4th subculture on DKW medium containing
1.0 µM of each 2, 4-D + TDZ (3.1 x).
119
6.2 CALLUS INDUCTION FROM IMMATURE FRUIT
The excised cotyledonary portions from immature fruits (Fig. 6.27) were cultured
on MS, DKW or WPM media containing different growth regulators (2, 4-D and TDZ).
Four different concentrations of 2, 4-D (4.52, 13.57, 22.61 or 31.65 µM) and three of
TDZ (10.0, 50 or 100 µM) were tested for callus induction (Table. 6.2).
BB
T
C A
T
B
F D E
Fig. 6.27: A) Immature Pecan (Carya illinoensis) fruits attached to a twig, collected
during August 2007. B) An opened view of immature fruit. C) Mature fruits collected
during September 2007, showing ruptured outer green husk. D) Mature fruits with a
pointed tip (T) and rounded base (B) (arrows) showing outer reddish-brown hard
endocarp with green husk removed. E) A longitudinal view of opened fruit from outside.
F) A longitudinal view of opened Pecan fruit cut from the centre into two halves (3.1 x).
120
TABLE: 6.2 EFFECT OF DKW, MS AND WPM MEDIUM WITH DIFFERENT LEVELS OF 2, 4-D ON CALLUS INDUCTION FROM IMMATURE FRUIT EXPLANTS
OF PECAN
Medium
Concentrations
(µM)
Callus induction (%) Morphological features
4.52 76.66 (11)* ± 8.81ab Yellowish-brown, friable, watery, translucent, loose
13.57 93.33 (11) ± 3.33a Yellowish-brown, compact, watery, translucent
22.61 76.66 (10) ± 8.81ab Off-White, friable, translucent at base
yellowish above, compact
DKW
31.65 70.00 (10) ± 5.77abc Yellowish-brown, compact and whitish-brown, friable, smooth
4.52 83.33 (11) ± 8.81 a Transparent-white, watery, loose, smooth
13.57 86.66 (12) ± 3.33a Translucent, watery, smooth, compact 22.61 73.33 (11) ± 3.33abc Brownish-translucent, watery, rough MS
31.65 53.33 (11) ± 8.81bcd Off-white, translucent, watery, friable, rough
4.52 50.00 (13) ± 5.77cd Brownish-white, watery, compact, rough
13.57 73.33 (15) ± 12.01abc Creamy-white, compact, granular 22.61 33.33 (15) ± 8.81cd Off-white, compact, smooth, watery WPM
31.65 46.66 (15) ± 3.33cd Whitish and brown, watery, compact, translucent
* The values in parenthesis in column for callus induction (%) are the number of days to callus induction. → Each value represents mean (± standard error) of three callus cultures and the experiment was repeated thrice → Values with different letters within a specific column represent significant difference at P<0.05 according to Duncan’s Multiple Range Test.
121
During the experimental period, immature fruit explants (cotyledonary parts)
cultured on all media with different levels of TDZ did not show any response in terms of
callus induction (Fig. 6.28 - 6.29). However, callus induction was observed on all the
tested levels of 2, 4-D on all media (DKW, MS, WPM) used. From all the media as
indicated in Table. 6.2, maximum callus induction (93.33 %) and proliferation was
observed on DKW medium containing 13.57 µM 2, 4-D after 10 days of initial culture.
Callus was yellowish-brown, translucent, watery and compact after which no further
response was observed (Fig. 6.30 - 6.32). This was followed by 76.66 % callus induction
on DKW medium supplemented with 4.52 and 22.61 µM 2, 4-D after 11 days of initial
culture (Table. 6.2).
Fig. 6.28 Fig. 6.29
Fig. 6.28: Immature Pecan fruit (cotyledonary portion) cultured on DKW medium
supplemented with 50 µM TDZ (3.1 x).
Fig. 6.29: A batch of culture vessels showing cultured fruit parts on MS medium
containing13.57 µM 2, 4-D (3.1 x).
122
Fig. 6.30 - 6.32: Callus morphology of Pecan fruit explants at 13.57 µM
2, 4-D supplemented to DKW medium at 25 ± 2 °C
Fig. 6.30 Fig. 6.31 Fig. 6.32
Fig. 6.30: Yellowish-brown, translucent, watery and compact callus formed on
DKW medium supplemented with 13.57 µM 2, 4-D at day 27 (2.0 x).
Fig. 6.31: Yellowish-brown, smooth and compact callus formed on DKW
medium supplemented with 13.57 µM 2, 4-D at day 39 (3.1 x).
Fig. 6.32: Yellowish-brown, compact and watery callus formed on DKW
medium supplemented with 13.57 µM 2, 4-D after 51 days (4.0 x).
Morphology of callus cultures formed on DKW medium containing 4.52 µM 2,
4-D is shown in Fig. 6.33 - 6.37. Two types of callus cultures, i.e., off-white, friable,
translucent and yellowish-brown, compact callus was observed on DKW medium
supplemented with 22.61 µM 2, 4-D (Fig. 6.36 - 6.37). Two types of calli were also
observed on DKW medium supplemented with 31.65 µM 2, 4-D. Calli were yellowish-
brown, compact, nodular and whitish-brown, compact, lustrous in appearance (Fig. 6.38-
6.39).
123
Fig. 6.33 - 6.35: Callus morphology of Pecan fruit explants at 4.52 µM 2, 4-D
supplemented to DKW medium at 25 ± 2 °C
Fig. 6.33 Fig. 6.34 Fig. 6.35
Fig. 6.33: Yellowish-brown, translucent, watery and compact callus formed on
DKW medium supplemented with 4.52 µM 2, 4-D (4.0 x).
Fig. 6.34: Yellowish-brown, smooth and compact callus formed on DKW
medium supplemented with 4.52 µM 2, 4-D (2.5 x).
Fig. 6.35: Translucent, granular, watery and compact callus formed on DKW
medium containing 4.52 µM 2, 4-D (4.0 x).
Fig. 6.36 - 6.37: Callus morphology of Pecan fruit explants at 22.61 µM
2, 4-D supplemented to DKW medium at 25 ± 2 °C
Fig. 6.36 Fig. 6.37
Fig. 6.36: Off-white, friable, translucent callus observed on DKW medium containing
22.61 µM 2, 4-D (4.0 x).
Fig. 6.37: Yellowish-brown, compact callus observed on DKW medium supplemented
with 22.61 µM 2, 4-D (3.1 x).
124
Fig. 6.38 - 6.39: Callus morphology of Pecan fruit explants at 31.65 µM
2, 4-D supplemented to DKW medium at 25 ± 2 °C
Fig. 6.38 Fig. 6.39
Fig. 6.38: Yellowish-brown, compact, nodular callus observed on DKW medium
supplemented with 31.65 µM 2, 4-D (2.0 x).
Fig. 6.39: Whitish-brown, compact, lustrous callus on DKW medium containing
31.65 µM 2, 4-D (3.1 x).
In MS medium containing 13.57 µM 2, 4-D, maximum (86.66 %) callus
induction was observed after 12 days of initial culture (Table. 6.2). Callus was
translucent, watery, smooth and compact (Fig. 6.40 - 6.41). A fair (83.33 %) callus
induction was observed on MS medium containing 4.52 µM 2, 4-D with transparent
white, watery, rough surface morphology (Fig. 6.42 - 6.43) followed by (73.33 %) light
brown, translucent, watery and smooth callus (Fig. 6.44 - 6.45) on DKW medium
supplemented with 22.61 µM 2, 4-D.
125
Fig. 6.40 - 6.41: Callus morphology of Pecan fruit explants at 13.57 µM
2, 4-D supplemented to MS medium at 25 ± 2 °C
Fig. 6.40 Fig. 6.41
Fig. 6.40 - 41: Translucent-white, watery callus also showing the greenish fruit
part (arrow) developed on MS medium containing 13.57 µM 2, 4-D
(1.7 x).
Fig. 6.42 - 6.43: Callus morphology of Pecan fruit explants at 4.52 µM 2, 4-D
supplemented to MS medium at 25 ± 2 °C
Fig. 6.42 Fig. 6.43
Fig. 6.42 - 43: Transparent white, watery, smooth callus highlighting the
whitish and green fruit part (arrows) developed on MS medium
containing 4.52 µM 2, 4-D (2.0 x).
126
Fig. 6.44 - 6.45: Callus morphology of Pecan fruit explants at 22.61 µM
2, 4-D supplemented to MS medium at 25 ± 2 °C
Fig. 6.44 Fig. 6.45
Fig. 6.44: Brownish-white, rough, compact callus on MS medium containing 22.61 µM
2, 4-D (3.1 x).
Fig. 6.45: Brown, smooth, compact callus highlighting the whitish fruit part
(arrow) developed on MS medium containing 22.61 µM 2, 4-D (2.0 x).
Fig. 6.46 - 6.47: Callus morphology of Pecan fruit explants at 31.65 µM
2, 4-D supplemented to DKW medium at 25 ± 2 °C
Fig. 6.46 Fig. 6.47
Fig. 6.46 - 47: Off-white, rough, compact callus highlighting the greenish-yellow
fruit part (arrow) developed on MS medium containing 31.65 µM 2, 4-D
(2.8 x).
127
WPM medium containing 13.57 µM 2, 4-D supported 73.33 % callus induction
after 15 days of initial culture (Table. 6.2). Callus was compact, creamy-white,
translucent and smooth (Fig. 6.48). Light yellowish-brown, translucent, watery and
smooth callus was observed on WPM medium supplemented with 4.52 µM 2, 4-D (Fig.
6.49). WPM medium containing 22.61 or 31.65 µM 2, 4-D, favored induction of off-
white, compact, granular or whitish-brown, watery, translucent callus, respectively, after
15 days of initial culture (Fig. 6.50 - 6.51). However, once the callus induction was
observed, further proliferation using these media was rather limited.
Fig. 6.48 - 6.49: Callus morphology of Pecan fruit explants at two levels of
2, 4-D (4.52 or 13.57 µM) supplemented to WPM medium at 25± 2 ºC
Fig. 6.48 Fig. 6.49
Fig. 6.48: Creamy-white, translucent, watery and smooth callus on WPM containing
13.57 µM 2, 4-D (3.1 x).
Fig. 6.49: Light yellowish-brown, watery and smooth callus observed on WPM
supplemented with 4.52 µM 2, 4-D (3.1 x).
128
Fig. 6.50 - 6.51: Callus morphology of Pecan fruit explants at two levels of
2, 4-D (22.61 or 31.65 µM) supplemented to WPM medium at 25± 2 ºC
Fig. 6.50 Fig. 6.51
Fig. 6.50: Off-white, granular, compact callus on WPM containing 22.61 µM
2, 4-D (3.1 x).
Fig. 6.51: Induction of light-brown, granular, watery callus (red arrow)
highlighting whitish fruit part (black arrow) on WPM containing 31.65
µM 2, 4-D (3.1 x).
Maximum (93.33 %) growth and proliferation of callus cultures was observed on
all the tested levels of 2, 4-D in DKW medium. Calluses obtained from all media (Table.
6.2) were maintained up to 2nd subculture by transferring them onto the respective fresh
medium. Further maintenance of such callus cultures was not possible due to sudden
browning (Fig. 6.52). Some of these cultures, however, were maintained for another 110
days using the same respective media. In such long-term callus cultures, initiations of
root primordia were observed (Fig. 6.52 - 6.56). This indicated a fair morphogenetical
potential of such callus cultures even though culture browning had already started.
Nonetheless, under the experimental conditions mentioned, shoot regeneration was not
obtained. Root initiation, similarly grew for a limited period of time. Such roots did not
show further response in terms of growth and development and became brown and
necrotic after 13 days (Fig. 6.57 - 6.59).
129
Fig. 6.52 - 6.56: Acute browning of callus developed on DKW medium (4.52
or 13.57 µM 2, 4-D) maintained at 25±2°C. Root initiation is also evident.
Fig. 6.52 Fig. 6.53 Fig. 6.54
Fig. 6.55 Fig.6.56
Fig. 6.52: Induction of root primordium (arrow) after callus browning on DKW
medium containing 4.52 µM 2, 4-D (3.1 x).
Fig. 6.53: Induction of root primordium (arrow) after callus browning on DKW
medium containing 13.57 µM 2, 4-D (2.1 x).
Fig. 6.54: Root initiation (arrows) from the callus (after callus browning had just
begun) developed on DKW medium containing 13.57 µM 2, 4-D
(3.1 x).
Fig. 6.55: An enlarged view of the Fig. 6.54 highlighting its left portion. Root
initiation (arrow) is quite evident (4.0 x)
Fig. 6.56: Right-side enlarged view from the Fig. 6.54 showing induction of two
roots (arrows) (3.1 x).
130
Fig. 6.57 - 6.59: Browning and necrosis of root primordia from fruit callus cultures developed on DKW medium containing 13.57 µM 2, 4-D at 25± 2 °C
Fig. 6.57 Fig.6.58 Fig. 6.59
Fig. 6.57: A developing root originating from a callus culture on DKW (4.52 µM
2, 4-D) showing signs of browning at day 13 of induction (4.0 x).
Fig. 6.58: Browning of root primordium (arrows) developed from a brown callus
at day 16 of induction (3.1 x).
Fig. 6.59: Completely necrotic callus at day 110 maintained on DKW medium
containing 13.57 µM 2, 4-D (4.0 x).
131
DISCUSSION
In vitro studies for Pecan (Carya illinoensis) improvement throughout the world
are generally scanty. Thus, there is a wide scope for further improvement of methods
ensuring callus induction and subsequent reproducible regeneration. In comparison with
herbaceous plants, lack of suitable explants presents a limiting factor to initiate in vitro
work in woody plants (Aftab et al., 2005). In addition, rejuvenation is another difficult
barrier in the regeneration of plants in such mature woody plants. It seems that tissues
that have undergone phase change are not easily converted back to juvenile
characteristics (Wareing, 1987). Hiatt and Allen (1991) state that “a plausible approach is
to use explant material that has not gone through phase change”. Owing to the above
knowledge and perhaps more specifically due to a hard woody texture of the bark, it has
never been used as an explant source in woody plant tissue culture. Prior to this study,
usually immature fruits (cotyledons/ zygotic embryos) were used for callus induction or
somatic embryogenesis (Merkle et al., 1987; Corte-Olivares et al., 1990b) in Pecan. In
addition, there is perhaps no report about the use of thidiazuron (N-phenyl-N'-1, 2, 3-
thidiazol-5-yl-urea; TDZ) in tissue culture studies of Pecan. TDZ exhibits a unique
property of mimicking both auxin and cytokinin effects on growth and differentiation of
cultured explants. It induces a diverse array of culture response ranging from induction
of callus to the formation of somatic embryos (Murthy et al., 1998). Its use in plant tissue
culture of recalcitrant woody plants has shown promise for micropropagation as well as
callus induction and regeneration studies (Huetteman and Preece, 1993; Wilhelm, 1999;
Preece et al., 2001; Preece, 2003; Ledbetter and Preece, 2004).
The mature bark explants were cultured on different media (DKW, MS or WPM)
supplemented with various levels of growth regulators (2, 4-D or TDZ) to investigate
callogenic response in Pecan. Although the bark explants, owing to their specific rough-
132
textured morphology could have been prone to contamination, the contamination rate
was surprisingly almost negligible in this work. It is evident form the results of the
present investigation that mature bark explants when cultured on DKW medium
containing a combination of 2, 4-D and TDZ supported 93.70 percent callus induction.
However, even with TDZ (50 µM) alone, 90.18 percent callus induction was recorded.
Similarly, WPM medium with 2, 4-D and TDZ (1µM each) produced 85.77 percent
callus followed by 65.33 percent on WPM containing 50 µM TDZ. Huetteman (1988)
reported that a higher concentration of TDZ (> 1.0 µM), could stimulate callus
formation, adventitious shoots or somatic embryos in J. nigra. On the other hand,
debarking (removal of bark) of trees during the vegetative period due to wounding
results in the formation of callus tissue which develops over the entire wound surface or
on parts of it (Stobbe et al., 2002). However, during the wood formation season, wounds
produced by complete removal of bark leaving the cambium or undifferentiated xylem
remains unaffected may lead to the formation of a callus tissue over the whole wound
surface (Kielbaso and Hart, 1997; Dujesiefken et al., 2001). Stobbe et al., (2002) in a
study involving lime trees revealed that, in most cases wounds separate the bark from
xylem within the zone of differentiating xylem while in some cases wounded surface
comprises of xylem, the innermost phloem or cambium cells alone. In some other
wounds incompletely differentiated xylem was exposed. They further supported the fact
that undifferentiated xylem cells at the stage of primary wall formation, proliferated
through mitotic activity, thus contributing to the callus formation. Formation of this type
of callus has been variously described as “reproduction of new bark and wood tissue”
(Hartig, 1844), “surface or superficial callus growth” (Sharples and Gunnery, 1933) or
just “surface callus” (Dujesiefken et al., 2001). During the present research work it was
also observed that callus was developed usually from the cut surfaces of the bark. This
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indicated that the bark explant must have partially differentiated xylem or actively-
dividing cambial cells that directed the formation of callus.
The present investigation also revealed that the season of bark collection
significantly influenced the callus induction. Bark explants collected during winter
season did not show any response in terms of callus induction. From spring-grown
(February - March) mature bark, 93.33 % callus induction was recorded on TDZ + NAA
(1.0 + 1.0 µM). Several factors have been reported to influence the activity of phellogen
layer of mature bark (Fahn, 1997; Stobbe et al., 2002). For instance, Stobbe et al., (2002)
reported that in lime trees (Tilia sp.) phellogen was found to be active during the
vegetative period (June). In Robinia, phellogen was found to be active mainly under a
combination of short day and high temperature (Borger and Kozlowski, 1972). The
results of the present investigation showed that a better callus induction response might
have been associated with an increased phellogen activity in spring season. Gibberellic
acid and naphthalene acetic acid were reported to have a retarding effect on phellogen
activity in Robinia (Borger and Kozlowski, 1972). As mentioned, TDZ, on the other
hand, is a potent cytokinin for woody plant tissue cultures (Huetteman and Preece,
1993). It might be possible that TDZ during this investigation enhanced the phellogen
activity. It was also evident that TDZ combined with 2, 4-D was more effective over 2,
4-D alone for the initiation of callus from immature bark explants of Pecan and its
subsequent maintenance. This investigation also suggested DKW medium to be the best
medium for in vitro callus induction and proliferation in Pecan (Carya illinoensis) as the
highest callusing percentage and proliferation was observed from the bark explants.
However, MS medium did not show satisfactory results in terms of callogenesis from
bark explants. Callus cultures were also transferred for plant regeneration. However,
plant regeneration was not possible during the present investigation.
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It is also evident from the results of the present study that cotyledonary portions
used as explants underwent 93.33, 86.66 or 73.33 percent callus induction at 13.57 µM 2,
4-D on the three tested media, i.e., DKW, MS or WPM respectively. As in bark explants,
callus was also developed from the cut surfaces of cotyledonary portions. It is revealed
from this study that DKW medium was most responsive and WPM found to be the least
responsive towards callus induction and proliferation. However, Long and coworkers
(1995) suggested that agar-solidified WPM supplemented with 0.1µM 2, 4-D and 50 µM
TDZ was the best treatment for the induction of somatic embryos and adventitious shoots
from immature cotyledonary explants of Juglans nigra L. (Eastern Black Walnut). Fruit
explants (cotyledonary portions) cultured during the months of September - October
have shown better response towards callus induction as compared to those cultured
during November- January. Similar findings were reported for Pecan (Wetzstein et al.,
1989) where several factors were shown to influence callus induction and induction
frequency of embryogenic cultures, i.e., effect of cultivars, sampling date, tree source of
explants and duration on conditioning medium in Pecan. In 1994, Rodriguez and
Wetzstein also investigated callus production, embryo formation and embryo
morphology in Pecan. They suggested that NAA at a concentration of 2, 6 or 12mg/litre
was a superior auxin over 2, 4-D for the callus proliferation and somatic embryos
development. It was revealed form results of the present study that callus cultures
obtained from DKW medium were maintained up to 2nd subculture, after which no
further proliferation was observed. In other media (MS or WPM), maximum callus
induction was recorded but no further development in terms of growth and proliferation
was observed. Callus cultures did tend to become brown and necrotic afterwards.
During the present work, it was observed that callus cultures obtained from bark
explants could be maintained successfully upto 4th subculture on DKW medium while
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callus cultures obtained from cotyledonary explants were maintained upto 2nd
subculture. Further maintenance of such callus cultures was not possible due to sudden
browning and blackening that was usually hard to control. Lee et al., (1990), reported
that this callus browning, necrosis and seizure of callus growth in many in vitro-grown
plants may be due to the accumulation of phenolic compounds and brown colour is due
to the formation of quinones which are inhibitory to callus growth. Browning of the
tissue is correlated with excessive accumulation of phenolics (Dubravina et al., 2005).
Ozyigit (2008) demonstrated that the darkening of cut or dying plant parts is caused by
the oxidation of phenolic acids and formation of polymers (dark aggregates). Ozyigit et
al., (2007) in a study involving Gossypium hirsutum, demonstrated the fact that no tissue
lacks phenolic compounds and high concentrations can be found in actively growing
cells. This phenomenon was also observed in some economically important plants, e.g.,
coffee, mango, chickpea, (Iqbal et al., 1991) guava, date palm, (Daayf et al., 2003) and
cotton (Ozyigit et al., 2007; Ozyigit, 2008). However, the callus cultures form the
present research work, after browning formed root-like structures kept on the same
medium for 110 days or so. No shoots were developed and these root primordia also did
not show any further development. This indeed highlighted the organogenetic potential
of such seemingly brown, necrotic callus cultures. The results of this investigation with
Pecan suggest control of browning to be of utmost importance. However, it also suggests
maintaining of even brown (though not necrotic?) callus cultures and necessitates further
attempts to figure out a possible plant regeneration protocol.
The aim of this study was to explore new means and response of even ‘atypical’
explants (mature bark, cotyledons) for callus induction and a reliable regeneration system
in Pecan through callus cultures. In conclusion, response of mature bark explants to
callus induction has shown promise since it has never been considered before a suitable
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explant source for callus induction or somatic embryogenesis due to its hard woody
texture. The results of this investigation have thus opened a possibility that such hard
bark portions might have a potential to dedifferentiate forming callus and hence, add to
the list of explants having potential for callus induction and somatic embryogenesis. It
also represents an ideal tissue for learning various aspects of in vitro growth and
differentiation in Pecan. Further studies incorporating other factors will probably lead to
a reproducible regeneration system in Carya from bark explants. TDZ as a growth
regulator has never been investigated in tissue culture of Pecan. The results of the present
study have proven TDZ to be a potent growth regulator for callus induction from bark
explants of Pecan.
It can be concluded that in vitro callus induction using bark as an explant source
is perhaps a newer approach in tissue culture studies of Pecan. The results of this
investigation demonstrated that bark can be a better explant source thus providing a rapid
method for callus induction. It also indicates a strong possibility to regenerate plants
using various tissues of Pecan. The results presented here seem encouraging and suggest
further studies on similar parameters so as to arrive at a meaningful, reproducible
protocol regarding in vitro Pecan cultures that may further be exploited in several studies
including genetic transformation of Pecan.
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CHAPTER 7
ADVENTITIOUS REGENERATION OF PECAN USING
IMMATURE COTYLEDONARY EXPLANTS
RESULTS
The present investigation demonstrates the effect of TDZ and BAP on the
development of adventitious shoots from immature cotyledonary explants in C.
illinoensis. The embryonic axes were excised carefully and small pieces of cotyledons
were removed and cultured on different media, i.e., DKW (Driver and Kuniyuki, 1984),
WPM [Woody Plant Medium (McCown and Lloyd, 1981)] or MS (Murashige and
Skoog, 1962) containing either BAP or TDZ at a concentration of 0.5, 1.0, 4.0, 8.0 or
15.0 µM.
The results from present investigation showed the induction of adventitious
multiple shoots from immature cotyledonary explants of Pecan (Carya illinoensis
(Wangenh.) C. Koch). Cotyledonary explants from Pecan seeds did not show any
response in terms of adventitious regeneration in WPM medium supplemented with any
tested level of BAP (Table. 7.1). The data presented in Table. 7.1 also reveal that
different levels of TDZ supplemented to all media (DKW, MS or WPM) also did not
show any response towards adventitious regeneration. Adventitious shoots were
developed from the cotyledonary explants of Pecan on MS or DKW medium
supplemented with 4.0, 8.0 or 15.0 µM BAP. In addition, adventitious shoots also
developed on MS medium supplemented with 1.0 µM BAP (Table. 7.1; Fig. 7.1).
However, MS medium supplemented with 15.0 µM BAP showed induction of maximum
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(12.1) multiple adventitious shoots followed by 8.0 µM BAP, after 15 days of initial
culture (Fig. 7.2 - 7.4). A bunch of multiple shoots were developed from cotyledonary
portions of fruit cultured on MS medium supplemented with 4.0 µM BAP (Fig. 7.5). A
few shoots were induced on MS medium supplemented with 1.0 µM BAP (Table. 7.1;
Fig. 7.6 - 7.7).
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TABLE: 7.1 EFFECT OF DIFFERENT LEVELS OF BAP OR TDZ SUPPLEMENTED TO
DKW, MS OR WPM MEDIUM ON ADVENTITIOUS SHOOT INDUCTION USING IMMATURE COTYLEDONARY EXPLANTS OF PECAN
PGRs Concentrations (µM)
Number of adventitious shoots/ explant
BAP
TDZ DKW MS
WPM
0.5 - NR NR NR 1.0 - NR 2.0 ± 1.10d NR 4.0 - 6.5 ± 0.25bc 4.1 ± 1.71cd NR 8.0 - 7.2 ± 1.02b 6.3 ± 0.98bc NR 15.0 - 10.9 ± 0.95a 12.1 ± 1.25a NR
- 0.5 NR NR NR - 1.0 NR NR NR - 4.0 NR NR NR - 8.0 NR NR NR - 15.0 NR NR NR
Means followed by same letters under different treatments within columns are not significantly
different using Duncan’s Multiple Range Test (P<0.05)
± represents standard error
NR: Not recorded
0
10
20
30
40
50
60
70
80
Shoo
t ind
uctio
n (%
)
0.5 1 4 8 15
BAP (uM)
MSWPMDKW
Fig. 7.1: Effect of different concentrations of BAP on in vitro shoot multiplication from
immature cotyledon of Pecan on three different salt formulations, i.e., DKW,
MS or WPM. Data were recorded at day 15 of initial culture.
140
EA
Fig. 7.2 Fig. 7.3 Fig. 7.4
Fig. 7.5 Fig. 7.6 Fig. 7.7
Fig. 7.2: Multiple shoots (right bracket) developed from immature cotyledonary portions
(arrow) on MS medium supplemented with 15.0 µM BAP (1.6 x).
Fig. 7.3: Multiple shoots (arrows) originating from immature embryonic axes (EA with
arrow) on MS medium supplemented with 15.0 µM BAP (1.2 x).
Fig. 7.4: Proliferating multiple shoots (right arc) on MS medium supplemented with 8.0
µM BAP (1.2 x).
Fig. 7.5: Multiple shoots (arrows) proliferating on MS medium supplemented with 4.0
µM BAP (1.3 x).
Fig. 7.6-7.7: Multiple shoot (arrows) induction on MS medium supplemented with 1.0
µM BAP at day14 of initial culture (2.5 x).
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On the other hand, maximum (10.9) number of shoots was developed on DKW
medium supplemented with 15.0 µM BAP (Fig. 7.8). A bunch of multiple shoots were
developed from cotyledonary portions of fruit cultured on DKW medium supplemented
with 8.0 µM BAP (Fig. 7.9 - 7.10). DKW medium supplemented with 4.0 µM BAP
showed the induction of an average (6.5) number of adventitious shoots (Table. 7.1; Fig.
7.11). However, no shoots were formed from the explants placed on WPM medium with
either BAP or TDZ (Table. 7.1; Fig. 7.1). The adventitious multiple shoots developed
from different media were further transferred to rooting media. Half of the proliferated
shoots (2.0 - 4.5 cm long) were transferred to MS basal medium and half were pre-
treated with IBA (1000 or 2000 ppm) for root induction. In either treatment, however,
rooting could not be induced even after 65 days of initial culture. Finally, the shoots
became necrotic.
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Fig. 7.8 Fig. 7.9
Fig. 7.8: Multiple shoots (left bracket) originating from immature cotyledonary portions
on DKW medium supplemented with 15.0 µM BAP (1.2 x).
Fig. 7.9: A bunch of multiple shoots (left bracket) originating from cotyledonary
portions on DKW medium supplemented with 8.0 µM BAP (3.1 x).
Fig. 7.10 Fig. 7.11
Fig. 7.10: Multiple shoots (arrows) proliferating on DKW medium containing 8.0 µM
BAP (1.3 x).
Fig. 7.11: Multiple shoots (arrows) originating from immature cotyledonary portions on
DKW medium supplemented with 4.0 µM BAP (1.3 x).
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DISCUSSION
Adventitious regeneration means the production of adventitious buds or shoots
from tissues other than the axillary buds, e.g., the cotyledonary explants. Although
adventitious regeneration is generally undesirable for clonal micropropagation, it
represents an excellent source of regenerated plants from various tissues. According to
Kantia and Kothari (2002), “adventitious organogenesis or shoot formation is a preferred
system as it enables to retain the clonal fidelity since many ornamental species are
cultivars that are propagated for one or more unique characteristics”. This phenomenon
may be of particular significance for extremely recalcitrant woody plant species. The
most common explant source for adventitious regeneration of woody plants is
cotyledons. They may either be from mature (Pooler and Scorza, 1995; Canli and Tian,
2008) or immature seeds/ cotyledons (Ainsley et al., 2001) or leaf tissues from in vitro
cultures (Messeuger et al., 1993; Kantia and Kothari, 2002). In vitro adventitious bud
regeneration has also been achieved from various explants of several woody tree species
including immature cotyledons of Juglans nigra (Long et al., 1995), epicotyl and
hypocotyl explants of Citrus sinensis (Maggon and Singh, 1996), hypocotyl segments of
Annona squamosa (Nagori and Purohit, 2004) and leaf and cotyledonary explants of
Crataegus pinnatifida (Dai et al., 2007). The propagation rates by means of shoot
organogenesis can be much higher than axillary shoot proliferation (Chun, 1993).
Adventitious shoot formation may also be used for overcoming reproductive barriers
caused by sterile male/female plants (Kantia and Kothari, 2002).
The role of BAP in adventitious shoot bud differentiation has been demonstrated
in a number of cases using variety of explants (Mao et al., 2000; Tawfik and Noga, 2001;
Nagori and Purohit, 2004; Purohit et al., 2004). It is evident from the present
experimental results that the highest number of shoots (12.1) per explant were produced
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after 16 days on MS medium supplemented with 15 µM BAP. MS medium
supplemented with 15 µM BAP was quite effective for 80 % shoot induction followed by
DKW (70 %) with the same BAP concentration. However, lower BAP concentrations
supplemented either to MS or DKW medium resulted in reduced shoot induction
potential. In another study, Obeidy and Smith (1993) showed adventitious buds arising in
callus cultures of mature Carya illinoensis embryonic tissue. Shoots were regenerated
from explants placed on MS (Murashige and Skoog, 1962) medium with 25 µM TDZ. In
another study involving Juglans nigra (another member of Juglandaceae), Long et al.,
(1995), has previously reported the formation of somatic embryos and adventitious
shoots from immature cotyledonary explants on WPM supplemented with 0.1 µM 2, 4-D
and 50 µM TDZ and greatest number of shoots per explant (28.9) were recorded. In
another research work on eastern black walnut (another nut- producing Pecan relative),
Neuman et al., (1993) did not observe the formation of adventitious shoots from the
immature cotyledonary explants on WPM with 2, 4-D and TDZ. Similarly, the results of
present investigation showed that no shoots were formed from the explants placed on
WPM medium with either BAP or TDZ.
Since, 1988, TDZ has been reported to induce adventitious shoot formation in a
number of species, especially woody plants (Lu, 1993). Although, considered to be most
effective in woody plant tissue culture (Huetteman and Preece, 1993), various levels of
TDZ supplemented to all three media, i.e., DKW, MS or WPM did not show any
response in terms of adventitious regeneration. In a study by Tang et al., (2002)
involving Prunus spp, TDZ could not initiate shoot bud differentiation. The present
study demonstrated BAP to be more effective and contributed towards 12.1 %
adventitious shoot development. Among the levels of BAP, 15.0 µM produced the
maximum number of shoots. There are prior reports highlighting the role of BAP in
145
comparison with TDZ in promoting shoot bud regeneration in various Prunus species
(Tang et al., 2002; Gentile et al., 2002). The results from the present investigation as
well as those mentioned above support each other and emphasize the fact of specific
requirements of growth regulators for a certain growth phenomenon. The results,
however, may not rule out the usefulness of a certain growth regulator which may be of
significance under a different experimental set-up or genotype.
The adventitious shoots developed in the present investigation showed a fair
response in terms of further growth and proliferation on transfer to MS basal medium.
Rooting of adventitious shoots, however, could not be induced even after 65 days of
initial culture. Rooting potential seems to be quite limited in this recalcitrant woody tree
species. In one such study, an average of 40 % rooting was observed after treatment of
adventitious shoots of Juglans nigra L. (Eastern Black Walnut) with 2.5 mM IBA and
1.25 mM NAA in dimethyl formamide (Long et al., 1995). Most woody plant species are
recalcitrant to adventitious organ formation (regeneration) due to genetically driven in
vitro recalcitrance (James et al., 1988; McCown, 2000; Singh et al., 2002). In addition,
fruit trees are amongst the most recalcitrant for in vitro culture, and regeneration of
adventitious shoots from adult explants has proven difficult (Miguel et al., 1996; Singh
and Sansavini, 1998). The results from this research work showed that Pecan is also
recalcitrant to adventitious regeneration, as the percentage of adventitious shoot buds
from cotyledonary explants was less than 50 %. Future work holds promise in enhancing
further proliferation and rooting potential of regenerated shoots thereby paving way for
successful establishment of plants under greenhouse conditions.
In conclusion, results from the present investigation demonstrate that adventitious
shoots can be developed successfully from the immature cotyledonary explants of Pecan.
The adventitious regeneration indicates a strong possibility to regenerate plants using
146
various tissues of Pecan. Although, rooting could not be induced during the present
investigation but due to the assumed comparative ease of the production of whole plants
(Long et al., 1995) this technique may possibly be advantageous over the other
conventional methods suggesting immature cotyledons as promising explants. Further
work holds promise in enhancing rooting potential and moving towards the development
of a reliable and reproducible method for Pecan regeneration by adventitious means.
147
CHAPTER 8
GENERAL DISCUSSION AND FUTURE WORK
The aim of the present research work was to propagate Pecan using tissue culture
procedures. Secondly, it also focused to explore response of various explants including
even ‘atypical’ ones (mature bark) for callus induction in an effort to develop a reliable
regeneration system in Pecan through callus cultures. Finally, its focus included
optimizing conditions for the establishment of forced softwood shoots for further
vegetative or micropropagation.
As a recalcitrant woody tree species, propagation of Pecan using tissue culture
techniques is quite a challenge. Conventionally, Pecan is propagated through seeds,
grafting and/or budding (Menary et al., 1975), however, these methods were
unsuccessful to raise large quantities of Pecan propagules. To accomplish the first goal
of this investigation, mature Pecan seeds were germinated in soil under glasshouse
conditions. The glasshouse conditions did not favor good germination response (merely
13.3 %). The presence of hard seed coat might well be a physical barrier that hampered
the germination of Pecan seeds under those conditions. Pecan seeds were therefore also
germinated under in vitro conditions on different media (DKW, MS or WPM) after
carefully removing the outer hard seed husk. The results proved MS basal medium to be
the most suitable one for in vitro seed germination. In addition, incision on the seeds
improved germination rate in Pecan. The percent seed germination response was also
remarkably enhanced on MS medium supplemented with BAP. Multiple shoots were
also observed to be developing from in vitro-germinating seedlings on media
supplemented with various levels of BAP. Highest number of multiple shoots (5.68) was
148
observed on DKW medium with 4.0 µM BAP. This work therefore has revealed a key
role played by BAP in seed germination in vitro as well as in the development of
multiple shoots. It further revealed a combination of IBA and NAA (4.0 + 4.0 µM)
supplemented to MS medium to be the best in terms of root induction in Pecan. With the
development of aforementioned protocol, more than 85 % of the plants were successfully
established in the field.
Mature bark and immature fruit (cotyledonary segments) were used as an explant
source and cultured on three different media (DKW, MS or WPM) containing various
levels of growth regulators (2, 4-D or TDZ) for callus induction. The results of the
present investigation have shown that a combination of 2, 4-D and TDZ (1.0 µM each)
brought about 93.70 % callus induction using mature bark on DKW medium, however, a
relatively much higher level of TDZ (50 µM) alone induced 90.18 % callus on DKW
medium. Callus formation was usually observed from the cut surfaces of the bark
signifying the fact that the bark explants had partially-differentiated xylem or actively-
dividing cambial cells. Explant collection during different seasons has also influenced
callus induction significantly. Winter season did not favor the formation of callus
whereas spring-grown mature bark explants resulted in 93.33 % callus induction on MS
medium supplemented with TDZ + NAA (1.0 µM each). The results of the present
research showed that a better callus induction response might have been associated with
an increased phellogen activity in the spring season. Additionally, during this
investigation TDZ might have enhanced the phellogen activity. On the other hand,
DKW medium favored highest (93.33 %) callus induction with 2, 4-D using
cotyledonary explants. DKW medium was found to be a better-supportive medium
towards callus induction and proliferation. Sudden blackening and necrosis though was a
major limitation in the maintenance of these callus cultures any longer. This callus
149
browning and arrest of callus growth might perhaps be due to an excessive accumulation
of phenolic compounds. However, such brown callus cultures from the present work
formed root-like structures after 110 days or so. No shoots were produced and the root
primordia also did not show any further growth. Nonetheless, this feature highlighted the
organogenetic potential of such apparently brown, necrotic callus cultures.
The present investigation also attempted to explore relatively newer and
alternative approaches for clonal propagation of Pecan. This aspect of the present work
has been accomplished through forcing shoot tips or large stem segments from the more
juvenile portions of the older trees during the dormant season. During the course of this
study, it was observed that forcing media (coccopeat, sand or sawdust) significantly
influenced the production of softwood shoots from large stem segments. The results
revealed that sterilized sand markedly affected the production and development of
softwood shoots. Maximum softwood shoots (2.92) were observed in sterilized sand.
Sand is a highly porous medium having excellent drainage properties, therefore tested as
growth medium because Pecan generally grows best on well-drained sandy loam or
loamy sand. Moreover, environmental conditions in lab, glasshouse or wire house also
had a significant effect on forcing softwood shoots from epicormic buds of large stem
segments of Pecan. The results of the present study revealed that glasshouse conditions
even without mist or fog system favored highest (2.92) mean number of softwood shoots.
The results were quite promising demonstrating an efficient and relatively economical
method for forcing softwood shoots in Pecan. Furthermore, production of softwood
shoots from large stem segments was also influenced by seasons. During the present
studies maximum numbers of softwood shoots was obtained during the winter season in
comparison to spring or autumn seasons. Stem diameter of logs also had a pronounced
effect on softwood shoot forcing in Pecan. The softwood shoots developing from stem
150
segments of diameter ≥2.5 cm have shown vigorous shoot development. During the
present research work softwood shoot forcing was also observed from stem segments of
greater than 5.6 cm diameter. Fishel et al., (2003) suggested that there may be no upper
limit to stem diameter for forcing because the lowest segments of main stems produced
the most shoots. This generalization seems to hold good for Pecan as well where smaller
diameter logs (< 2.5 cm) showed reduced shoot forcing. Softwood shoots (≥ 4 cm)
obtained from large stem segments were subjected to rooting media (sand, peat moss,
perlite or vermiculite), however rooting was not achieved in the present set of
experiments.
During the present study, adventitious multiple shoots were produced
successfully on DKW or MS medium containing 4.0, 8.0 or 15.0 µM BAP from
immature cotyledonary explants of Pecan. The highest (80 %) number of shoots
originated on MS medium containing 15.0 µM BAP. Lower BAP concentrations,
however only showed reduced shoot induction potential. On the contrary, no shoot
formation was observed on WPM medium with either BAP or TDZ. Even various levels
of TDZ did not favor any adventitious regeneration when supplemented to all three
tested media, i.e., DKW, MS or WPM. The results of the present study have thus shown
BAP to be more effective over TDZ in adventitious shoot development. Among the
levels of BAP, 15.0 µM produced the maximum number of shoots. The results from the
present investigation emphasize the need of a specific growth regulator for a certain
growth phenomenon. Once again the results of the present investigation revealed rooting
of the adventitious shoots to be a major constraint in Pecan. The present investigation demonstrates that due to several limitations in
conventional breeding procedures, in vitro seed germination not only enables the
production of aseptic seedlings in short duration but also useful for the large scale
151
production of Pecan root stock. In vitro-raised seedlings may also be of advantage where
genetic variability is desirable. Further studies in this regard may play an important role
in the multiplication and improvement of this recalcitrant tree species. Callus induction
from bark explants in this study seems encouraging and suggests further investigation on
similar parameters so as to arrive at a meaningful, reproducible protocol regarding
development of in vitro Pecan cultures. Such in vitro cultures may then further be
exploited in several studies including genetic transformation where availability of a
suitable plant material is still considered to be a major bottleneck. During this
investigation, the possibility of forcing epicormic buds from large stem segments of
Pecan has opened up a way leading to the establishment of a cost-efficient method to
produce a sufficient number of Pecan shoots that may be used directly for clonal
propagation or to be exploited for micropropagation. The results from this study
highlighted a fact that even much cheaper and easily procurable media such as silica sand
could satisfactorily be used for softwood shoot production in Pecan though other tested
media also had a potential to promote satisfactory results in this regard. Further testing of
silica sand and other media as used in this study for softwood shoot initiation is therefore
recommended for this and other temperate or tropical woody tree species as well.
The present study demonstrates suitability of novel propagation methods to be
exploited in Pecan. Thus softwood shoot forcing was successful along with the
establishment of other protocols for in vitro seed germination, micropropagation, callus
induction and further differentiation, and adventitious shoot regeneration in Pecan.
Suitability of mature bark explants for callus induction and somatic embryogenesis adds
it to the potential list of explant sources. Immature cotyledons have also shown
promising results for the development of a consistent and reproducible method for
adventitious regeneration. Rooting of softwood or adventitious shoots although seems to
152
be a challenging aspect for Pecan propagation using these means and thus could only
partially be accomplished during the present study. In conclusion this study has opened
up a direction for further investigation in Pecan. By no means its benefits for a larger
scale Pecan propagation may be harnessed directly unless and until the bottlenecks
mentioned above and throughout this study are further worked upon and remaining gaps
carefully worked out and patched-up. Potential use of the mentioned protocols and
methods though is highly promising. It is hence quite likely in the future that further
work from this and other labs may overcome current limitations to harness the full
potential that these protocols and methods hold for Pecan improvement.
153
CHAPTER 9
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180
ANNEXURE-I
FORMULATION OF DKW MEDIUM (DRIVER AND KUNIYUKI, 1984) FOR
PREPARATION OF STOCK SOLUTION
Constituents
Stocks Concentration in DKW
Medium A. ORGANICS (DKW1 )
Myo-inositol Glycine Nicotinic acid Thiamine-HCl
mg/ l (50x) 100 × 50 = 5000 100 50 100
mg/ l 100 2.0 1.0 2.0
B. PHOSPHATES (DKW2)
KH2PO4 H3BO3 Na2MoO4.2H2O
mg/ l (50x) 264.8 × 50 =13240
240 19.5
mg/ l 264.8 4.8 0.39
C. NITRATES (DKW3)
NH4NO3 Ca(NO3)2 Zn(NO3)2
mg/ l (50x) 1416 × 50 = 70800
98400 850
mg/ l 1416
1968 17
D. CALCIUM (DKW4) CaCl2
mg/ l (50x) 149 × 50 = 7450
mg/ l 149
E. SULPHATES (DKW5)
MgSO4.7H2O MnSO4.4H2O
CuSO2.5H2O K2SO4
mg/ l (50x) 1560 × 50 = 78000
3700 1675
12.5
mg/ l 1560 740 33.5
0.25
F. IRON EDTA (DKW6) Na2-EDTA FeSO4
mg/ l (50x) 45.4 × 50 = 2270
1690
mg/ l 45.4 33.8
181
ANNEXURE-II
FORMULATION OF MS MEDIUM (MURASHIGE AND SKOOG, 1962) FOR PREPARATION OF STOCK SOLUTION
Constituents Stocks Concentration in MS Medium
A. MACRONUTRIENTSNH4NO3 KNO3 CaCl2.2H2O MgSO4.6H2O KH2PO4
mg/ l (20x) 1650 × 20 = 33,000 38,000 8,800 7,400 3,400
mg/ l 1,650 1,900 440 370 170
B. MICRONUTRIENTS
MnSO4.4H2O ZnSO4.4H2O H3BO3 KI Na2MoO4.2H2O CuSO4.5H2O CoCl2.6H2O
mg/ l (100x) 22.3 × 100 = 2,230 860 620 83 25
2.50 2.50
mg/ l 22.3 8.6 6.2 0.83 0.25 0.025 0.025
C. IRON-EDTA
Na2-EDTA.2H2O FeSO4.7H2O
mg/ l (200x) 27.8 × 200 = 5,560
7,440
mg/ l 27.8 37.2
D. VITAMINSGlycine Nicotinic acid Pyridoxine-HCl Thiamine-HCl
mg/ l (100x) 2.0 × 100 = 200 100 100 20
mg/ l 2.0 0.5 0.5 0.1
E. MYO-INOSITOL
mg/ l (100x)
100 × 100 = 10000
mg/ l
100
182
ANNEXURE-III
FORMULATION OF WPM MEDIUM (MCCOWN AND LLOYD, 1981) FOR
PREPARATION OF STOCK SOLUTION
Constituents Stocks Concentration in WPM Medium
A. MACRONUTRIENTS NH4NO3 K2SO4 CaCl2.2H2O MgSO4.7H2O KH2PO4 Ca(NO3)2 .4H2O
mg/ l (20x) 400 × 20 = 8000 19800 1920 7400 3400 11120
mg/ l 400
990 96 370 170 556
B. MICRONUTRIENTS H3BO3.7H2O MnSO4.4H2O ZnSO4.7H2O CuSO2.5H2O Na2MoO4.2H2O
mg/ l (100x) 6.2 × 100 = 620
2230 860 25 25
mg/ l 6.2 22.3 8.6 0.25 0.25
C. IRON-EDTA Na2EDTA.2H2O FeSO4.7H2O
mg/ l (20x) 37.2 × 20 = 744 556
mg/ l 37.2 27.8
D. VITAMINS Glycine Nicotinic acid Pyridoxine-HCl Thiamine-HCl
mg/ l (100x) 2 × 100 = 200 50 50 100
mg/ l 2.0 0.5 0.5 1.0
E. MYO-INOSITOL
mg/ l (50x) 100 × 50 = 500
mg/ l 100
183
ANNEXURE-IV
GROWTH REGULATORS USED IN THIS STUDY WITH RESPECTIVE ABBREVIATION,
MOLECULAR WEIGHT AND INITIAL SOLVENT
Growth regulators
Abbreviation
Mol. weight (g)
Dissolve in*
2, 4 -dichlorophenoxy
acetic acid
2, 4 -D
221
1N NaOH/
Ethanol
Indole-3-acetic acid
IAA
175.19
1N NaOH/
Ethanol
Indole-3-butyric acid
IBA
203.24
1N NaOH/
Ethanol
6-benzylaminopurire
BAP
225.3
1N NaOH
Naphthalene Acetic
Acid
NAA
186.2
1N NaOH
Thidiazuron (N-phenyl-Nَ-1, 2, 3-
thiadiazol-5-yl urea)
TDZ
220.2
0.1N KOH
* This is the initial solvent to get the respective growth regulators dissolved. The final volume was made up by slowly adding distilled water to an appropriate level.
184
ANNEXURE-V Preparation of 1 liter DKW Medium
One liter DKW medium was prepared in a manner given below.
Medium Components Stock Concentration Volume of Stock solution
1) Organics 50X 20 ml/ l
2) Nitrates 50X 20 ml/ l
3) Sulphates 50X 20 ml /l
4) Calcium 50X 20 ml/ l
5) Phosphates 50X 20 ml/ l
6) Iron 50X 20 ml/ l
7) Sucrose 30 g/ l
8) Agar (Oxoid, Hampshire, England) 7.0 g/ l
9) pH 5.75 - 5.85
10) Growth regulator (2, 4-D, IAA, IBA, BAP, NAA, TDZ): According to the
requirement of a specific medium.
ANNEXURE-VI
Preparation of 1 liter MS Medium
One liter MS basal medium was prepared in a manner given below.
Medium Components Stock Concentration Volume of Stock solution
1) Macronutrients 20X 50 ml/ l
2) Micronutrients 100X 10 ml/ l
3) Vitamins 100X 10 ml/ l
4) Myo-inositol 100X 10 ml/ l
5) Iron-EDTA 200X 05 ml/ l
6) Sucrose 30 g/ l
7) Agar (Oxoid, Hampshire, England) 7.0 g/ l
8) pH 5.75 - 5.85
9) Growth regulators (2, 4-D, IAA, IBA, BAP, NAA, TDZ): According to the
requirement of a specific medium.
185
ANNEXURE-VII
Preparation of 1 liter WPM Medium
One liter WPM medium was prepared in a manner given below.
Medium Components Stock Concentration Volume of Stock solution
1) Macronutrients 20X 50 ml/ l
2) Micronutrients 100X 10 ml/ l
3) Vitamins 100X 10 ml/ l
4) Myo-inositol 50X 20 ml/ l
5) Iron-EDTA 20X 50 ml/ l
6) Sucrose 20 g/ l
7) Agar (Oxoid, Hampshire, England) 6.0 g/ l
8) pH 5.75 - 5.85
9) Growth regulators (2, 4-D, IAA, IBA, BAP, NAA, TDZ): According to the
requirement of a specific medium.
ANNEXURE-VIII
Composition of Different Media Used for In Vitro Germination of Pecan Seeds
Basal medium PGRs Concentrations
0.5 µM BAP
1.0 µM BAP
3.0 µM BAP
4.0 µM BAP
8.0 µM BAP
12.0 µM BAP
DKW, MS or WPM basal
or supplemented with
PGRs
15.0 µM BAP
186
ANNEXURE-IX
Composition of Different Media Used for Callus induction/Maintenance from
Mature Bark
Basal Medium Growth Regulators Concentrations
4.52 µM 13.57 µM 2, 4-D 22.61 µM 1.0 µM 50 µM TDZ
100.0 µM
DKW, MS or
WPM
TDZ + NAA 1.0 + 1.0 µM
ANNEXURE-X Composition of Different Media Used for Plant Regeneration from Callus Cultures
Medium Medium Composition
2.22 µM BAP
2.22 µM BAP + 0.5 µM TDZ MS
2.22 µM BAP + 1.0 µM TDZ
187
ANNEXURE-XI
Composition of Different Media Used for Callus induction/Maintenance from Immature/ Mature Fruits
Basal medium PGRs Concentrations
4.52 µM 2, 4-D
13.57 µM 2, 4-D DKW, MS or
WPM 22.61 µM 2, 4-D
ANNEXURE-XII
Composition of Different Media Used for Adventitious Regeneration of Pecan
Basal medium PGRs Concentrations
0.5 µM BAP or TDZ
1.0 µM BAP or TDZ
4.0 µM BAP or TDZ
8.0 µM BAP or TDZ
MS, WPM or DKW
15.0 µM BAP or TDZ