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

133

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.

134

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