UNDERSTANDING THE FLORAL TRANSITION IN AQUILEGIA COERULEA AND DEVELOPMENT OF A TISSUE CULTURE PROTOCOL

A Thesis

Presented to the

Faculty of

California State Polytechnic University, Pomona

In Partial Fulfillment

Of the Requirements for the Degree

Master of Science

In

Plant Science

By

Timothy A. Batz

2018

SIGNATURE PAGE

THESIS: UNDERSTANDING THE FLORAL TRANSITION IN AQUILEGIA COERULEA AND DEVELOPMENT OF A TISSUE CULTURE PROTOCOL

AUTHOR: Timothy A. Batz

DATE SUBMITTED: Summer 2018

College of Agriculture

Dr. Bharti Sharma Thesis Committee Co-Chair Department of Biological Sciences

Dr. Valerie Mellano Thesis Committee Co-Chair Plant Science Department

Dr. Kristin Bozak Department of Biological Sciences

ii ACKNOWLEDGEMENTS

I would like to thank the many faculty, family, and friends who helped me enormously throughout my master’s program. The endless support, mentorship, and motivation was crucial to my success now and in the future. Thank you!

Dr. Mellano, as my academic advisor and mentor since my freshman year at Cal Poly Pomona, I greatly appreciate your time and dedication to my success. Thank you for guiding me towards my career in science.

Dr. Sharma, thank you for taking me into your lab and taking the role of research mentor. Your letters of support allowed me the opportunities to grow as a scientist.

Dr. Bozak, I always had a pleasure meeting with you for advice and constructive critiques. Thank you for the time spent reading my statements and the opportunities to gain presentation skills by lecturing in your classes.

Dr. Still, thank you for introducing me into the world of research. Thank you for helping me understand the work and input required for scientific success.

To lab mates Summer Blanco, Jesus Preciado, and Michael Speck, thank you for helping make research enjoyable. It was a pleasure to learn and grow as scientists with you all!

To Dr. Adler, Dr. Buckley, Dr. Valdez, and Airan Jansen of the Cal Poly RISE Program, thank you for instilling values of hard work, determination, and cooperation in us budding scientists. It was pleasure learning from your experiences in science.

To Dr. Washburn, Dr. Columbus, and Dylan Cohen of the Rancho Santa Ana Botanic Garden, my work relied heavily on your diligent training and generous allowance of the facilities and instruments. Thank you all for your support and collaboration.

A mi familia amorosa y apoyoso, gracias por su amor y sacrificio. Eso me motivó a perseguir mis objetivos. Les dedico este trabajo a todos ustedes.

iii

ABSTRACT

Vernalization, or exposure to prolonged cold, is a trigger which initiates the transition to flowering in many plants including crop and ornamental species. This environmental change in temperature is prevalent at high altitude environments and has become a strong selection pressure influencing the reproductive biology of plants.

Species in the genus Aquilegia are primarily found in mountainous areas of Europe, Asia, and North America. In this study, we are using A. coerulea as a model system to understand the morphological and developmental changes that the shoot apical meristem undergoes during the transitioning from a vegetative to reproductive meristem. Plants were grown to maturity and then vernalized at 6°C for 4 weeks, then subsequently transferred to greenhouse conditions at 24°C. Meristems were dissected at each week throughout the treatments and prepared for histology and scanning electron microscopy.

Results indicate that the formation of the reproductive inflorescence begins at the third week of vernalization. Floral meristems are established by the fourth week of greenhouse conditions. All vernalized plants flowered around the fourth week of greenhouse conditions while nonvernalized plants failed to do so. We conclude that vernalization is essential for flowering in A. coeruela.

The ability to insert genes into plant models in vitro is a powerful tool for genetic investigations into morphological traits and developmental processes. A tissue culture protocol to establish transgenic plants is required. Seeds of A. coerulea were germinated in vitro and used as sterile explant tissues. Cotyledons grown in medium containing a

[1.0/1.0mgL-1] ratio of auxin and cytokinin plant hormones, BAP/2,4-D (6-

iv Benzylaminopurine/2,4-Dichlorophenoxyacetic acid) produced reliable callus growth.

Shoot proliferation from callus was promoted by a BAP/2,4-D ratio of [0.5/0.25mgL-1]. A liquid medium containing [1.0mgL-1] of the auxin IAA (Indole-3-acetic acid) was found to promote root organogenesis from shooted callus in combination with 7 days of dark treatment. With further optimization of media, a tissue culture protocol can be established to genetically transform and regenerate A. coerulea plantlets in vitro, a technique useful in forward and reverse genetics experiments performed in the fields of plant evolution and developmental biology.

v TABLE OF CONTENTS

Signature Page…………..….……………………………………………………………ii

Acknowledgements ...... iii

Abstract ...... iv

List Of Tables ...... viii

List Of Figures...... x

Abbreviations………………………………………..………………………………….xii

Chapter 1 Introduction...... 1

Background………………………………………………………………………..1

Literature Review………………………………………………………………….3

Chapter 2 Morphological & Developmental Studies To Understand The Floral

Transition In Aquilegia coerulea...... 11

Background………………………………………………………………………11

Objectives & Hypotheses……………………………………………………...... 14

Materials & Methods…………………………………………………………….15

Results……………………………………………………………………………30

Conclusions...…………………………………………………………………….43

Chapter 3 Tissue culture of Aquilegia coerulea ...... 44

Background………………………………………………………………………44

Objectives & Hypotheses………………………………………………………...45

Materials & Methods…………………………………………………………….46

vi Results…………………………………………………………………………....58

Conclusions...…………………………………………………………………….79

References……………………………………………………………………………….82

vii LIST OF TABLES

CHAPTER 2: MORPHOLOGY OF THE FLORAL TRANSITION

Table 1. Treatment of plants and number of meristems collected for histological analysis.

...... 17

Table 2. Tissue preparation protocol for histology ...... 19

Table 3. Staining procedure using Sharman series ...... 20-21

Table 4. Timepoints and conditions of plant collection and meristem dissection for SEM analysis before, during, and after vernalization treatment ...... 23

Table 5. Dehydration timetable for dissected meristems in preparation for SEM ...... 26

CHAPTER 3: IN VITRO REGENERATION OF AQUILEGIA COERULEA

Table 6. Solutions for sterilization of A. coerulea seeds ...... 47

Table 7. A. coerulea germination media: Ingredients and their respective amounts ...... 48

Table 8. Ratios and concentrations of BAP and 2,4-D used in callogenesis of A. coeruela cotyledons...... 51

Table 9. Concentration of AgNO3 used in callus media ...... 52

Table 10. Combinations of BAP, 2,4-D, and AgNO3 for callus induction of A. coerulea cotyledons ...... 53

Table 11. Combinations of BAP, 2,4-D, used for shoot and root induction in A. coerulea callus tissues...... 55

Table 12. Treatments of BAP, 2,4-D, and darkness (aluminum foil) used for root induction in A. coerulea shooted callus ...... 56

viii Table 13. Ratios of BAP/2,4-D medias A-I and their resulting percentage of induction and organogenesis in A. coerulea cotyledons after 6 weeks of culture ...... 63

Table 14. Media treatments with their respective ratios of BAP, 2,4-D, and AgNO3 ..... 67

Table 15. Ratios of BAP/2,4-D in callus medias and their resulting percentage of shoot and root induction in A. coerulea cotyledons after 9 weeks of culture ...... 75

Table 16. Number of roots produced from A. coerulea shooted calluses in

BAP/2,4-D and IAA liquid media ± 7 days of darkness ...... 77

ix LIST OF FIGURES

CHAPTER 2: MORPHOLOGY OF THE FLORAL TRANSITION

Figure 1. Approximate weekly growth rate of A. coerulea seedlings at 24°C...... 16

Figure 2. Sharman stain series solutions ...... 22

Figure 3. Stereoscopic dissection of A. coerulea meristems...... 24

Figure 4. Meristems of A. coerulea in vials of formaldehyde-acetic acid alcohol (FAA).

...... 25

Figure 5. Mounting of critical point dried A. coerulea meristems ...... 27

Figure 6. Sputter coating of dissected A. coerulea meristems in preparation for

SEM analysis ...... 28

Figure 7. Loading of sputter coated A. coeruela meristems into the scanning electron microscope chamber...... 29

Figure 8. Dissected shoot apical meristems of A. coerulea before, during, and after vernalization compared to the nonvernalized control ...... 30-31

Figure 9. Longitudinal sections of A. coerulea before, during, and after vernalization at 6°C compared to the nonvernalized control at 24°C ...... 32-33

Figure 10. Plan and side view orientations of A. coerulea meristems under scanning electron microscopy ...... 34-37

Figure 11. Comparison of vernalized and nonvernalized control plants at 4 weeks post-vernalization at 24°C ...... 37

x CHAPTER 3: IN VITRO REGENERATION OF AQUILEGIA COERULEA

Figure 12. Sterilization process used for handling all explant and cultured tissues ...... 47

Figure 13. Diagram of A. coerulea seed sterilization protocol in 6 steps ...... 48

Figure 14. Dissection of A. coerulea seedlings 2 weeks after germination ...... 50

Figure 15. Protocol for callus culture from A. coerulea seedling cotyledons in 4 steps...51

Figure 16. Glass vessels used for root and shoot cultures of A. coerulea callus ...... 54

Figure 17. Paper bridge cultures of A. coerulea callus using liquid media...... 56

Figure 18. Liquid culture of A. coerulea callus with shoots using a filter paper bridge method ...... 57

Figure 19. Dissected hypocotyl of A. coerulea seedling at week 0 and week 4 of callus culture in [0.5/0.5mgL-1] BAP/2,4-D...... 59

Figure 20. Growth of A. coerulea cotyledon callus tissues over 6 weeks of culture in BAP/2,4-D media...... 60-61

Figure 21. Rooting, shooting, and callus producing media before and after 6 weeks of tissue culture...... 64

Figure 22. Growth of A. coerulea cotyledon callus tissues over 6 weeks of culture in BAP/2,4-D ± AgNO3 media...... 65-66

Figure 23. Cotyledons of A. coerulea before and after 6 weeks of culture in

-1 -1 [1/1mgL ] and [2/2mgL ] ratios of BAP, 2,4-D ± AgNO3...... 68-71

Figure 24. Shoot organogenesis from A. coerulea callus tissues in [0.5/0.25mgL-

1] BAP/2,4-D after 9 weeks of culture in a 16L/8D photoperiod at 25°C...... 76

Figure 25. Adventitious root organogenesis from A. coerulea shooted calluses in

[1.0mgL-1] IAA + 7 days of dark treatment...... 77

xi ABBREVIATIONS

2,4-D……………………………………………………...2,4-Dichlorophenoxyacetic acid

#L/#D……………………………………………..(hours) Light/(hours) Dark photoperiod

AM……………………………………………………………………....Axillary meristem

BAP……………………………………………………………….....6-Benzylaminopurine

Bt………………………………………………………………………………...……Bract diH2O...…………………………………………………………………...Deionized water

IAA……………………………………………………………………Indole-3-acetic acid

IM………………………………………………………………….Inflorescence meristem

LFM………………………………………………………………..Lateral floral meristem

LP….……………………………………………………………………….Leaf primordia

MS…...……………………….Murashige and Skoog basal salt with vitamins and sucrose

Sp……...…………………………………………………………………..Sepal primordia

St…...……………………………………………………………………Stamen primordia

TB…………………………………………………………………………….Terminal bud

TFM…………...... ……….……………………………………...Terminal floral meristem

VM……………………………………………………………………Vegetative meristem

xii CHAPTER 1

INTRODUCTION

Background

The transition to flowering is a critical phase in the lifecycle of angiosperms. This is characterized by the production and formation of a single flower or many flowers clustered together in an inflorescence. This shift in development occurs when environmental and endogenous signals promote the expression of key floral genes. This switch from vegetative to reproductive growth is initiated in the shoot apical meristem.

Seasonal oscillations of temperature and daylength (photoperiod) greatly impact flowering time, along with other factors such as nutrient availability (Rolland et al.,

2006). Developmental changes and hormonal pathways in some plants ensure that flowering takes place regardless of the outside environment (Amasino & Michaels,

2010). The interactions of these internal, external, biotic, and abiotic factors act to maximize the reproductive success of the plant by initiating flowering when conditions are most conducive to fertilization and seed formation (Coupland, 1995).

Despite rapid advances in plant biology over recent decades, limitations in the knowledge of flowering time control exist. The plant kingdom encompasses rich variation in molecular, biochemical and developmental adaptations to aerial, aquatic, and terrestrial environments. This is demonstrated in the reproductive biology of the currently known 350,000 flowering plants. The morphological and molecular mechanisms behind the flowering transition are only well understood in a few species (Mandoli & Olmstead,

2000). Foundational studies on the flowering transition have been based on established plant models like Arabidopsis, Maize (corn), and Oryza sativa (rice). However, one

1 model system cannot represent the diversity of physiological, molecular, and environmental factors involved in flowering across the angiosperms (Mandoli &

Olmstead, 2000).

The development of powerful genetic tools and in vitro regeneration methods are expanding the number of plant model systems available for study (Mandoli & Olmstead,

2000). The genus Aquilegia (columbine), is an emerging model system suited for answering key questions related to floral morphology, development, evolution, and genetics (Kramer, 2009). Few studies have been done using this model, and most with a focus on evolution and ecology (Hodges & Kramer, 2007; Kramer, 2009; Kramer &

Hodges, 2010, Sharma et al., 2011).

Ballerini and Kramer (2011) investigated the reproductive development of

Aquilegia formosa and revealed that vernalization, or prolonged cold exposure of 6-8 weeks, is a requirement necessary for the induction of flowering in that species (Ballerini

& Kramer, 2011). Studies in Arabidopsis and O. sativa suggest that vernalization pathways have evolved independently in different plant species (Alexandre & Hennig,

2008). Compared to the lifecycle of A. Formosa and other related species in the genus, A. coerulea is rapid cycling. This study aims to understand the stage of development at which the shoot apical meristem transitions to an inflorescence meristem, and whether this process occurs during the relatively shorter 4 week vernalization period or after plants are removed from cold treatment.

2

Literature Review

Photoperiod and flowering time

The decision to flower occurs as a response from the integration of environmental and endogenous signals sensed by the plant. Due to the static nature of plants, they have adapted to sense their spatial-temporal surroundings through their leaves by specialized pigments located in plastids of the leaf cells (Raven, 2012). The specialized structures are adapted to not only capture sunlight for photosynthesis, but to also measure the amount of available sunlight in a day.

Photoperiodism, the physiological response of an organism to available light and dark in a 24-hour period, is a main environmental cue involved in the flowering transition

(Klejnot & Lin, 2004). Annual rhythm of longer day lengths in spring/summer and shortening days in fall/winter is a reliable signal for the sensing of seasons, especially for plants adapted to temperate environments at higher latitudes (Markovskaya & Sysoeva,

2011). Depending on geographical location and physiological requirements, plants utilize photoperiod as one of the environmental cues to determine the initiation of flowering

(Amasino, 2010).

There is variation in this response to day length as not all plants flower in response to longer days. Short day (SD) plants transition to flowering in response to a short day and long night. Conversely, long day (LD) plants are induced to flower when exposed to long days and short nights (Jordan, 2006). Interestingly, Ballerini and Kramer

(2011) showed that photoperiodism alone has little to no effect on the induction of flowering in A. formosa.

3 Physiological experiments conducted in the 1930s showed that leaves are the major sensors of daylength (Knott, 1934). Knott defoliated the majority of spinach leaves and exposed the last mature leaf to favorable photoperiods. He found that spinach plants exposed to the inductive photoperiods flowered despite heavy defoliation. Control plants without any leaves failed to flower under the same conditions (Knott, 1934). Plants with intact foliage also flowered in inductive photoperiods, thus demonstrating that a single leaf is sufficient to promote flowering under appropriate photoperiodic conditions (Knott,

1934).

Work by Lang and colleagues revealed that a mobile flower-promoting hormone named “florigen” produced in the leaves is responsible for inducing flowering under inductive photoperiods required for a particular species. They experimented with grafts of tobacco plants “Trapezond” (day neutral), “Maryland Mammoth” (short-day), and

“silvestris” (long-day). Unions of day neutral and short-day plants in short-day conditions exhibited an accelerated flowering effect. The same results occurred in grafts of day neutral and long-day plants in long day photoperiods. When short-day/day neutral grafts were exposed to long days, flowering was slightly delayed compared to non-grafted, day neutral controls (Lang et al. 1977). Grafts of long-day/day neutral plants in short day photoperiods was inhibited and growth exhibited a dwarf-like condition. The study concluded that florigen is transmitted from the long or short-day grafts to the day neutral graft partner, and that it is vital in floral promotion in photosensitive plant species (Lang et al. 1977).

Although the floral transition in many biennial plants is characterized by a cold requirement, vernalization itself does not necessarily cause flowering. Rather, it renders

4 the plant competent to do so (Lang, 1965). Daylength, or photoperiod also has strong effect on transition to flowering. Prior work by Lang and Melchers (1965) studied the biennial Black Henbane (Hyoscyamus niger) which requires vernalization followed by inductive photoperiods to flower. They vernalized several Henbane plants and subsequently exposed them to non-inductive photoperiods where they continued vegetative growth. However, when the plants were placed in inductive photoperiods, they flowered (Lang, 1965). This experiment demonstrated that vernalized plants are able to

“remember” their past cold conditions. They had acquired flowering competency but did not actually flower until the correct photoperiod was met.

Mechanisms of the photoperiod pathway

Plants perceive light through photoreceptors, which include red/far-red light sensing phytochromes (phy) and blue/UV light sensing cryptochromes (cry). As the seasons progress, the quality of light changes and the amount of red/far-red and blue/UV light sensed by these photoreceptors fluctuates (Lin, 2000). The annual rhythm of these light wavelengths is “memorized” by the circadian clock, a regular oscillation of biological activity in a 24-hour cycle.

Coupland et al. discovered the gene CONSTANS (CO) in Arabidopsis that encodes a transcription factor critical to photoperiodic flowering (Putterill et al., 1995).

The CO protein activates genes required for floral initiation, including FLOWERING

LOCUS T (FT) (Samach et. al, 2000). Valverde et al. found that the CO protein is ubiquitinated, or inactivated, and then degraded by the proteasome protein complex. This process is regulated by both cryptochromes and phytochromes. Transgenic Arabidopsis constitutively expressing CO independent of circadian control showed that CO protein

5 levels were higher in the light phase of long days than in the light phase of short days, regardless of constant mRNA expression (Valverde et al., 2004).

Vernalization and the flowering response

Vernalization, or the acquisition of flowering competence by cold exposure

(Chouard, 1960), is another environmental cue vital to the reproductive response in temperate plants like Aquilegia. Whereas photoperiod is useful for sensing the changing of seasons through the length of day and night cycles, a decrease in average temperature provides plants with the cue of winter. Flowering in plants adapted to cold weather occurs after the passing of winter when warmer temperatures return in spring (Fernando

& Coupland, 2012). Vernalization leads the epigenetic suppression of floral repressors like FLC. This repression enables the expression of genes that promote flowering

(Amasino, 2005).

Angiosperm diversification over ~140 million years resulted in a convergent evolution of vernalization pathways in crucifers (Arabidopsis), Amaranthaceae (sugar beet) and Pooideae (wheat and barley) (Ream et al., 2012). Research over recent years in established models like Arabidopsis and O. sativa has only provided limited understanding of these genetic programs involved in vernalization (Ream et al., 2012).

More studies are needed in plants representing various groups such as the basal eudicots

The transition to flowering in Aquilegia will not occur without vernalization. Aquilegia species therefore serve as excellent models to study developmental and genetic effects of vernalization on flowering.

6 Vernalization flowering pathway

Research into the effects of cold temperatures on flowering began mainly in agricultural crops (Chouard, 1960). The first systematic study into temperature effects on flowering was done by German plant physiologist Gustav Gassner. His work showed the great variety of cold requirements across the plant kingdom. Gassner vernalized experimental plants in the dark, dismissing the need for artificial daylength conditions during vernalization. He was then able to match vernalized plants to nonvernalized control plants at the same developmental stage for comparison.

By measuring minimal chilling time in many species, Gassner was able to differentiate the specific requirements between winter and summer plants. He reported that some plants require cold treatment whereas others have no significant chilling requirement for flowering. Gassner also showed that in winter cereals, the early swollen germinating seed is already sensitive to the specific cold effect (Gassner, 1918).

Prolonged cold treatment of seeds promotes flowering of many temperate cereals, including wheat, barley, oat, and rye. When grown without cold treatment, these plants remain vegetative for long periods or do not flower at all. Thus, Gassner stated that these cereal varieties have a cold exposure requirement, or ‘Kaltbedurfnis’, which is key for their transition to reproductive growth (Gassner, 1918).

Further work on cold-induced flowering in grains was conducted by the infamous

Soviet agrobiologist Trofim Lysenko. In his 1928 paper, he treated winter wheat seeds with moisture and cold and induced them to flower when planted in spring, ahead of their usual fall planting. Lysenko described that cold-requiring biennial plants like winter wheat can be made to flower in one growing season by providing low temperature

7 treatment to young plants or moistened seeds. He called the effect of chilling treatment

“vernalization”, after the Latin word for spring, “vernum”. He therefore proposed that vernalization is a process of shortening the juvenile or vegetative growth phase and hastening flowering by a previous cold treatment (Lysenko, 1928). However, Lysenko wrongly asserted that the vernalized state could be inherited and that the offspring of a vernalized plant would no longer require cold treatment to flower. His ideas of improved heritability led to his involvement with the Stalin regime and the misuse of science with political ideology (Caspari & Marshak, 1965).

Although early investigations on vernalization were focused on plant physiology, later developments in genetics and molecular biology led to increased knowledge of underlying mechanisms involved in cold-induced flowering. In 1904, Correns investigated the genetic distinctions between summer-annual and vernalization-requiring biennial plants. He studied two forms (annual and biennial) of the same species, Black

Henbane (Hyoscyamus niger). He germinated seeds collected from annual and biennial forms in the same environment and found that annuals always produced annual offspring and biennials always produced biennial offspring. By crossing the two forms he discovered the F1 offspring were biennial. The F2 offspring showed segregation, suggesting that a single dominant gene determined the biennial habit (Correns, 1904). In the coming decades, the molecular revolution would uncover and characterize this gene and many others involved in the vernalization floral transition.

Mechanisms of the vernalization pathway

The establishment of Arabidopsis thaliana (thale cress) as a model plant species in the 1960’s led to a greater focus on the little known molecular aspects of plant biology.

8 The diverse forms of Arabidopsis, including rapid-flowering annual and biennial accessions, rapidly progressed the genetic understanding of vernalization mechanisms. In the same vein as Correns, Napp-Zinn crossed several biennial and rapid-flowering

Arabidopsis accessions and analyzed segregation in the offspring. He showed that the delayed flowering of the vernalization-responsive biennials is due to a dominant gene which be named FRIGIDA (FRI) (Napp-Zinn, 1987). This discovery lead to the identification of other dominant FRI alleles among the diverse Arabidopsis accessions.

Koornneef and colleagues further identified additional genes involved in vernalization in Arabidopsis. A mutant screen was conducted on methanesulphonate or irradiation treated plants under long day conditions. A total of 42 mutants were identified with mutations at 11 loci. The mutations were classified as recessive, intermediate, or dominant and were located in positions on 4 of the 5 Arabidopsis chromosomes. These mutants flowered later or as late as the late-flowering parental mutant and differed in vernalization response compared to the wild type. For mutants at the loci fca, fve, fy and fpa, vernalization has a large effect both under long day and short-day conditions.

Conversely, co, gi, fd and fwa mutants were almost completely insensitive to this treatment (Koornneef et al., 1991). Further work on these mutants continues to reveal genetic mechanisms involved in the floral transition under vernalization.

9 Studies in Aquilegia

Much of what is known about vernalization and flowering comes from the model plant Arabidopsis (Napp-Zinn, 1987), core eudicots like pea (Wellensiek, 1973), and monocots including rice and wheat (Yan, 2004). Similarities to vernalization have been found across related plant families through inherited homologous genes. However, genetic duplication and speciation events, in addition to spontaneous mutations and domestication result in a wide variation of genes involved in floral induction through vernalization. Studying floral induction through vernalization in Aquilegia, a basal eudicot, will provide an intermediate point of comparison between the two main eudicot and monocot plant groups (Ballerini and Kramer, 2011).

To date, few studies have been published regarding floral transition and development in Aquilegia. Most studies have focused on a particular species in the

Aquilegia genus, A. formosa. Unlike some Arabidopsis accessions, in which vernalization renders flowering competency, Ballerini and Kramer (2011) have identified vernalization as the inductive signal for flowering in A. formosa. When exposed to temperatures of

4°C, A. formosa meristems begin to develop a terminal bud at the SAM. During the 6th week of cold temperatures, two lateral buds form along an elongating axis, which eventually develops into an inflorescence after 8 weeks of vernalization (Ballerini and

Kramer, 2011).

10 CHAPTER 2

MORPHOLOGICAL & DEVELOPMENTAL STUDIES TO UNDERSTAND THE

FLORAL TRANSITION IN AQUILEGIA COERULEA

Background

The genus Aquilegia (columbine) (Ranunculaceae, Ranunculales) is an emerging model system in plant biology. It is of particular interest to the fields of evolution, ecology and development (Hodges & Kramer, 2007; Kramer, 2009; Kramer & Hodges,

2010). The unique floral morphology of columbines has been the subject of comparative histological, genetic and genomic studies investigating floral spur anatomy and the evolution of petal spur length (Antoń & Kamińska, 2015; Puzey et al., 2011, Sharma et al., 2011). The sequenced genome of Aquilegia coerulea has allowed for genetic investigations into these aspects of floral development and ecology, as well as evolutionary relationships to other species within the genus and across the plant kingdom

(Kramer et al., 2007; Whittall & Hodges, 2007).

The natural habitats of A. coerulea lie at high elevations (6,900 to 12,100 ft) within the Rocky Mountains of North America which experience seasonal oscillations of warm and cold temperatures (USDA, n.d.). At these extreme conditions, short growing seasons and limited pollinators have resulted in the evolution of strategies crucial for the reproductive success of plants (Kö rner, 2014). Vernalization, or exposure to prolonged cold, is an epigenetic cue that initiates the floral transition in many winter annuals and biennials (Amasimo, 2004). By sensing the passing of winter, plants can initiate flowering when conditions are conducive to flower development and pollination

11 (Amasimo, 2004). For plants that require vernalization, flower buds will not form until the cold requirement is fulfilled (Finical, 1998).

Previous studies on the floral development of Aquilegia have noted the importance of vernalization on its reproductive growth (Shedron & Weiler, 1982; White et al., 1990; Finical, 1998; Tucker & Hodges, 2005; Ballerni & Kramer, 2011). In 1982,

Shedron and Weiler tested the flowering response to vernalization in the A. coerulea variety “McKana’s Giant”. Plants grown to 16 weeks of maturity were vernalized at

4.5°C. Flowering responses differed with the length of cold treatment, with 50% of plants flowering after 8 weeks and 100% flowering after 10 weeks (Shedron & Weiler, 1982).

In 1998, Finical repeated the Shedron and Weiler experiment with “McKana’s

Giant” columbines, but although the plants were grown to the age 16 weeks from seed as specified by Shedron and Weiler, none of the plants flowered after cold treatments at 5°C

(Finical, 1998). Both studies by Shedron, Weiler, and Finical focused on the macromorphological level of flowering, or the emergence of the inflorescence stem and subsequent anthesis, which seemed to differ in response to vernalization (Shedron &

Weiler, 1982; Finical, 1998).

White et al. (1990) utilized scanning electron microscopy to study floral initiation and development in 13 Aquilegia cultivars. Floral initiation occurred without vernalization in the cultivars ‘Bluebird’, ‘Dove’, ‘Purple’, and ‘Robin’ 5 months after sowing. In the remaining cultivars, four initiation stages were observed: a vegetative state in which leaf primordia were formed in whorls around the SAM, the development of a terminal bud from the vegetative SAM, the formation of a three-parted compound cyme

12 inflorescence, and the formation of additional three-parted compound cymes produced in several lateral inflorescences (White et al., 1990).

In 2005, Tucker and Hodges used SEM analysis to study the floral ontogeny of the Aquilegia, Semiaquilegia, and Enemion genera of the family Ranunculaceae.

A spurred Aquilegia species, A. olympica, was studied throughout its development. The vegetative shoot tips converted into a flower subtended by bracteoles. Axillary flowers were borne terminally on an axis that also bore a bracteole pair. A total of 10 stamen orthostichies in the floral apices of A. olympica were observed (Tucker & Hodges, 2005).

In 2011, Ballerini and Kramer studied the floral transition in Aquilegia formosa and discovered a strong vernalization requirement, but little flowering response to changes by photoperiod alone. Mature plants were vernalized for eight weeks at 4°C in short day conditions (8L/16D) and then returned to either short day or long day (16L/8D) greenhouse conditions. Short day plants flowered 29 days post-vernalization and long day plants after 20 days (Ballerini & Kramer, 2011). To test whether a shift in daylength influenced the flowering response, long day plants were not vernalized and maintained at

20°C. Of the nonvernalized plants, 78% failed to flower, emphasizing the importance of cold treatment in the floral transition in A. formosa (Ballerini & Kramer, 2011).

To understand when the meristem transitions into an inflorescence in response to vernalization, Ballerini and Kramer conducted histological studies of the apical meristems before vernalization, two weeks, six weeks, and eight weeks into vernalization, and one week post-vernalization at 4°C. Morphological analysis showed a cymose inflorescence in which the meristem transforms into a terminal bud that produces two lateral buds; the lateral buds repeat this pattern (Ballerini & Kramer, 2011). By the

13 first week of post-vernalization, the terminal bud produces stamen primordia and sepals which is mimicked by the axillary floral meristems. While photoperiod may have some impact on growth rate after vernalization, Ballerini and Kramer found that the meristem actually transitions to flowering during the cold treatment at 4°C (Ballerini & Kramer,

2011).

Objectives & Hypotheses

Decades of published studies have confirmed the shoot apical meristem as the main site of environmental signal perception and aboveground plant organogenesis

(Machida, Fukaki, & Araki, 2013). Leaves, shoots, and flowers all originate from the collection of stem cells at the shoot apex (Raven et. al, 2012). This region was studied in

A. formosa by Ballerini and Kramer in 2011. Using histology and staining techniques, the transition of the shoot apex from a vegetative to inflorescence meristem was observed under vernalization treatment at 4°C for 8 weeks (Ballerini & Kramer, 2011). According to the Syngenta Aquilegia coerulea “Origami” culture guide, vernalization for this species is best achieved when night temperatures reach a range of 5-10°C for a period of

14-21 days, with average post-vernalization daily temperatures between 18°-19°C

(Syngenta, n.d.). With this information, we hypothesize that the reproductive transition in

A. coerulea will occur in the shoot apical meristem at some point during the vernalization period of 4 weeks at 6°C or in subsequent greenhouse conditions between 18-19°C.

14 Materials & Methods

Cultivation and vernalization of Aquilegia coerulea

Columbine seeds of the Aquilegia coerulea variety, “Origami Red and White”

(swallowtailgardenseeds.com), were sowed on a perforated seed tray with PRO-MIX BX

Mycorrhizae soil mix (Premier Tech Horticulture). After germination, 3-inch-tall seedlings were transplanted into 4.5-inch pots containing PRO-MIX and Nutri-Rich 4-3-2

Fertilizer Pellets. Plants were watered only when the soil was dry to avoid root rot. The presence of white fly, caterpillars and Downy mildew on the plants was continuously monitored and regular removal of brown and dried leaves was conducted to minimize pests. Plants were grown in a greenhouse without supplemental lighting with an average day and night temperatures at 24°C and 18°C, respectively. Plant growth was steady in greenhouse conditions, with each plant producing an average of one leaf per week at

24°C (Figure 1). Out of the 68 plants observed, most reached the 8-leaf stage (indicating maturity) after 8-9 weeks of cultivation and were ready for vernalization treatment.

To observe developmental changes in the shoot apical meristem (SAM) during the reproductive transition, plants were grown to the 8-leaf stage in 4-inch pots and vernalized (Figure 1). A growth chamber (Percival E41VL, USA) was utilized to simulate winter conditions. A total of 50 A. coerulea plants were vernalized. Growth conditions in the chamber were set to a 16-hr day/8-hr night photoperiod with 60% humidity, 80% light intensity, and a constant temperature of 6°C for 4 weeks. This temperature was chosen based on optimum range for effective vernalization which is near or at 6°C (Street & Öpik, 1984). A Syngenta flower culture guide for Aquilegia coerulea recommended a temperature range within 5°-10°C for effective vernalization (Syngenta,

15 n.d.). Several culture guides of other Aquilegia species recommended chilling at a narrower degree of 5°C for Auilegia x hybrida “Origami Blue and White” and Aquilegia vulgaris (Whitman & Runkle, 2012; Whitman, & Padhye, 2008).

Figure 1: Approximate weekly growth rate of A. coerulea seedlings at 24°C. Leaf number from 68 plants was counted over 10 weeks to determine the point of maturation. Plants were responsive to vernalization treatment at the 8-leaf stage denoted by the orange line.

Vernalized plants were subsequently moved into greenhouse conditions of approximately 24°C for an additional 4 weeks. Meristem collections occurred at weekly points before, during, and after vernalization treatment. Meristems of pre-vernalization and nonvernalized control plants were also dissected for comparison (Table 1). A group of five plants was collected simultaneously to check for developmental coherence in the meristems at that particular time point. The shoot apex was dissected and the surrounding

16 rosette of leaf tissue was removed with forceps. The meristems were fixed in formalin– propionic acid–alcohol (FPA) immediately following dissection.

Table 1: Treatment of plants and number of meristems collected for histological analysis. Samples of the shoot apical meristem were dissected with razor and forceps and prepared for histology via fixation, dehydration, and wax infiltration steps described in table 3.

Treatment Week of experiment # of samples Environmental temp. Pre-vernalization 0 5 ~ 24°C Vernalization 1 5 6°C Vernalization 2 5 6°C Vernalization 3 5 6°C Vernalization 4 5 6°C Post-vernalization 5 5 ~ 24°C Post-vernalization 6 5 ~ 24°C Post-vernalization 7 5 ~ 24°C Post-vernalization 5 ~ 24°C 8 Non-vernalized control 5 ~ 24°C

Histology

For histological studies, the morphological state of the collected meristem samples was preserved from dehydration and decay through a biological fixation.

Samples were placed in vials of formalin–propionic acid–alcohol (FPA), vacuum infiltrated to remove air from within the tissues, and left overnight in FPA. Following this, the meristems progressed through a dehydration series of increasing ethanol solutions to draw out all water from within the tissues (70, 90, 95, 100 w/safranin, and

100%) (Table 2). Sample colors fade due to the increasing ethanol treatments, leaving the small samples colorless. To improve sample visibility for downstream manipulation, a

17 staining step of 100% EtOH with 1% w/v safranin was included during the dehydration series. Dehydration procedures ended with meristems in 100% ethanol.

Following dehydration, the meristems underwent a ‘clearing’ step in which the ethanol dehydrant within the intercellular spaces was replaced with the miscible solvents, xylene and paraffin oil (Ruzin, 1999). To displace the ethanol in the meristems, dehydrated samples were immersed into several ratios of decreasing ethanol, increasing xylene (2:1 and 1:2 ratios of 100% EtOH:Xylene), and pure xylene (Table 2). The xylene in the tissues was then replaced with paraffin oil through several solutions of increasing paraffin concentration (2:1 and 1:2 ratios of Xylene:Paraffin oil). A minimum soaking time of 2 hours was required for the dehydration, clearing, and wax infiltration steps. The final infiltration steps required soaking the meristems in two solutions of paraffin wax at

6 hours per change (Table 2).

After wax infiltration, the meristems were embedded in paraffin wax using a histoembedder (Leica EG1160, Germany). Samples were carefully oriented within a

7x7x5mm plastic mold (HistoPrep cat. no. 15-182-501A) during contact with the melted paraffin wax. The position of the meristem was stabilized using forceps until the mold was filled with paraffin. Sample orientation was crucial to achieving a longitudinal view of the meristems during sectioning. A plastic cassette (Ted Pella INC. cat. no. 27159-3) was subsequently placed on top of the mold containing the sample and covered with additional paraffin to fuse the mold and cassette together. After wax embedding, the meristems were left to cool slowly at room temperature until the melted paraffin solidified.

18 Table 2: Tissue preparation protocol for histology. Samples of the shoot apical meristem were dissected with razor and forceps and prepared for histology via fixation, dehydration, clearing, and wax infiltration prior to embedding, sectioning and staining.

Step Solution Time Tissue Fixation FPA (vacuum infiltration) Overnight 70% EtOH 2 hours 90% EtOH 2 hours Tissue 95% EtOH 2 hours Dehydration 100% EtOH 2 hours 100% EtOH + 1% w/v Safranin Overnight 100% EtOH 2 hours 2:1 100% EtOH:Xylene 2 hours 1:2 100% EtOH:Xylene 2 hours Tissue 100% Xylene 2 hours Clearing 100% Xylene 2 hours 2:1 Xylene:Parrafin oil 2 hours 1:2 Xylene:Parrafin oil 2 hours Wax Paraffin (1st change) 6 hours Infiltration Paraffin (2nd change) 6 hours

Meristem samples were sectioned using a microtome (Leica RM 2135, Germany).

The blade was set at a 10° angle, resulting in sections 10um thick. The wax ribbons containing the longitudinal sections of the meristems were trimmed to form sections 5cm in length and laid on top of a 1.2mm thick glass slide (Gold Seal cat. no. 3048) covered with Sass’s Adhesive Solution (Ruzin, 1999) and diH2O. After, the slides were placed on a slide warmer (Model 77, Fisher Scientific) set at 42°C to heat the water and allow full expansion of the wax ribbons. After ribbon expansion, the slides were cleared of excess diH2O and left to dry in a warming oven overnight at 42°C.

19 After the slides were completely dried, they were stained for tissue visibility using the Sharman stain series (Sharman, 1943) commonly used in histological analysis of shoot apex tissues. Aqueous solutions of diluted 2% ZnCl2, 1% iron alum, 5% tannic acid

+ Orange g, and safranin were used to stain cell walls, starch grains, nuclei, and lignified cell walls respectively (Figure 2) (Ruzin, 1999). CitriSolv was used to dissolve paraffin and expose the tissues to the staining agents. Solutions were prepared in volumes of

500ml and contained in staining dishes. Dried slides were placed in a slide holder and progressively moved from one dish to the other in the steps outlined in table 4. The slides remained in xylene (step 30) while they were removed one by one for coverslipping. To permanently seal the stained slides, a few drops of CytosealTM 60 (Thermo Scientific) were placed on top of the tissues. A 24x60mm (Fisher Scientific cat. no. 12-545-89) coverslip was dipped in xylene and placed over the glass slide to prevent drying of the stained tissues. The coverslipped, permanently sealed slides were then allowed to dry overnight.

Table 3: Staining procedure using Sharman series. After drying overnight, sectioned tissues on the glass slides were placed in a slide rack and moved from progressively from step 1 to 30 for staining. Steps 28-30 were done under a fume hood to reduce chemical exposure.

Step Solution Time (min) 1 CitriSolv 10 2 CitriSolv 10 3 1:1 100% EtOH: CitriSolv 5 4 100% EtOH 5 5 95% EtOH 2

Table continued on page 21.

20 Table 3: Staining procedure using Sharman series (continued).

Step Solution Time (min) 6 90% EtOH 2 7 70% EtOH 2 8 50% EtOH 2 9 30% EtOH 2

10 diH2O 2

11 2% aqueous ZnCl2 1

12 diH2O 5 seconds 13 1:25,000 aqueous safranin 5

14 diH2O 5 seconds 15 Orange G + tannic acid 1

16 diH2O 5 seconds 17 5% Tannic acid 5

18 diH2O 1-3 seconds 19 1% aqueous iron alum 2

20 diH2O 15 seconds 21 30% EtOH 5 seconds 22 50% EtOH 5 seconds 23 70% EtOH 5 seconds 24 90% EtOH 5 seconds 25 95% EtOH 5 seconds 26 100% EtOH 5 seconds 27 100% EtOH 10 seconds 28 (in fume hood) 3:1 xylene:methyl salicylate 2 29 (in fume hood) Xylene 2 30 (in fume hood) Xylene Remain until coverslipped

21

Figure 2: Sharman stain series solutions. Glass staining dishes were filled with 500ml of zinc chloride, iron alum, tannic acid, orange G and safranin to stain meristematic tissues to stain cell walls, starch grains, nuclei, and lignified cell walls, respectively (Sharman, 1943).

To observe changes in the meristems of A. coerulea throughout pre, mid, and post-vernalization treatments, microscopic analysis of prepared slides was conducted using a Laborlux D compound microscope (Leitz Wetzlar, Germany). Images were captured using the Olympus DP73 microscope camera attachment (Leitz Wetzlar,

Germany). Longitudinal sections of meristems of before, during, and after vernalization at 6°C and subsequent greenhouse treatment at 24°C were compared to a nonvernalized group of control plants to observe morphological changes in the shoot apical meristem

(Figure 9). Replicates of five A. coerulea plants were prepared and observed to confirm developmental synchrony across the vernalization and greenhouse treatments.

22 Scanning electron microscopy

Light microscope observations of the internal anatomy in 2-D were obtained from histology. To understand the 3-D morphological changes in the shoot apical meristems of

A. coerula under vernalization, scanning electron microscopy was employed. Following the treatment procedures from the histology experiment, 6 plants were dissected from each treatment group before, during, and after vernalization at 6°C to confirm a cohesive developmental shift in morphology (Table 4). An increased sample size for weeks 3 and

4 was designed to confirm the initiation of the floral transition as seen from the histology results (Figure 9 D, E). At each timepoint (Table 4), plants were removed from their pots, washed, and defoliated in preparation for meristem dissection. Using a stereomicroscope

(Leica EZ4D, Germany), mature and developing leaves were removed to reveal the shoot apical meristem (Figure 8 A-J).

Table 4: Timepoints and conditions of plant collection and meristem dissection for SEM analysis before, during, and after vernalization treatment. Increased number of meristems were dissected at weeks 3 and 4 to observe if the developmental was synchronous across samples.

Treatment Week of experiment # of samples Condition of samples Pre-vernalization 0 6 ~ 24°C Vernalization 1 6 6°C Vernalization 2 10 6°C Vernalization 3 10 6°C Vernalization 4 6 6°C Post-vernalization 5 6 ~ 24°C Post-vernalization 6 6 ~ 24°C Post-vernalization 7 6 ~ 24°C Post-vernalization 6 ~ 24°C 8 Nonvernalized control 6 ~ 24°C

23 A. B.

C. D.

Figure 3: Stereoscopic dissection of A. coerulea meristems. At each timepoint, plants were removed from soil (A) and the root crown and stem regions excised from the roots (B). The Leica EZ4D dissecting scope with a camera attachment was used for delicate removal of the leaf rosettes surrounding the shoot apical meristem (D).

24 Following dissection, meristems were chemically fixed in vials of formaldehyde- acetic acid alcohol (FAA) to prevent biological degradation of the tissues (Figure 4).

Once the samples from all 9 timepoints were collected (Table 4), the meristems were dehydrated in ethanol solutions of increasing concentration (Table 5) in preparation for the critical point drying process. Samples fully immersed in 100% EtOH were subsequently transferred to porous specimen holders under 100% EtOH conditions to avoid drying of the samples (Figure 5 A). These holders were then loaded into the critical point dryer chamber.

In preparation for SEM analysis, the critical point drying technique was used to delicately desiccate the samples prior to their introduction into the SEM vacuum chamber. This step was necessary to preserve the morphological features of the meristems which would otherwise be obliterated as the water inside the samples violently disassociates inside the vacuum chamber.

Figure 4: Meristems of A. coerulea in vials of formaldehyde-acetic acid alcohol (FAA). To preserve their biological state, dissected meristems were preserved in FAA (10% FAA; 50% pure EtOH; 5% glacial acetic acid; 35% H2O) and stored at 6°C.

25 Table 5: Dehydration timetable for dissected meristems in preparation for SEM. Following FAA fixation, A. coerulea meristems were dehydrated in 6 steps of increasing ethanol concentrations. The samples were immersed in two separate solutions of 100% EtOH to completely remove water from the samples.

Step EtOH solution Time (hrs) 1 70% 2 2 80% 2 3 90% 2 4 95% 2 5 100% 2 6 100% 2

The ethanol within the samples was replaced by the transfer fluid (liquid CO2) by controlled release from a storage tank connected to the critical point chamber. This process was augmented by simultaneously cooling the chamber to 20°C through control of the cold-water valve connected to the chamber and the nearby sink. The amount of liquid CO2 was kept above the sample holders by balancing the CO2 tank and the vent valves to avoid backpressure. Once the meniscus of the liquid the CO2 stabilized above the samples in the chamber, a 3-4 minute flushing step was conducted to fully remove the ethanol substitution liquid from the samples. To do so, the inlet valve was fully opened, the vent valve was closed, and the drain valve was opened to release the ethanol-liquid

CO2 mixture from the chamber. After flushing, the samples were left inside the liquid

CO2 filled chamber for a 1-hour impregnation step.

Following impregnation, the chamber was once more flushed for 3-4 minutes with liquid CO2 by opening the inlet and vent valves while maintaining the meniscus above the samples. To raise the liquid CO2 above its critical point of 31.1°C, all valves to the

26 chamber were closed and the chamber temperature was slowly increased to 36°C via the faucet. At this point, the meniscus observed in the chamber was no longer visible, indicating that the intermediate phase between a liquid to gas (critical point) has been reached. To avoid condensation of the transfer liquid and specimen damage caused by rapid venting, the CO2 was slowly released over 3-4 minutes by opening the vent valve.

The dried specimens were removed from the chamber and mounted on stubs in preparation for sputter coating (Figure 5 C, D).

A. B.

C. D.

Figure 5: Mounting of critical point dried A. coerulea meristems. After ethanol dehydration, samples were placed in porous holders (A) and desiccated through critical point drying. The samples were then carefully oriented in plan and side views on stubs (B, D) in preparation for sputter coating (C).

27

A. B.

C.

Figure 6: Sputter coating of dissected A. coerulea meristems in preparation for SEM analysis. Samples were coated using the 108auto (A) under vacuum conditions with a gold target (B). A 15nm thick gold layer was applied to all meristems (C).

The meristems were positioned in plan and side view orientations for full coverage of morphological features (Figure 5 B-D). Sputter coating was done using the

108 Auto Sputter Coater (Ted Pella, INC., Redding, CA) with a gold target. The samples were loaded onto the rotating stage chamber which was then flushed with argon gas to create an inert environment conducive to unobstructed coating. A gold layer 15nm thick was formed on all samples for conduction of the electron beam inside the SEM chamber

(Figure 6 B, C). The gold coated meristems were then loaded into the SEM vacuum chamber (Hitachi SU3500, Hitachi High Technologies America, Inc.) and observed filament settings at 10-15 kV (figure 7A, B). SEM images were taken at several magnifications per sample and analyzed using MountainsMap® Universal software

(Digital Surf, France).

28

A. B.

Figure 7: Loading of sputter coated A. coeruela meristems into the scanning electron microscope chamber. Gold coated samples (B) placed into the Hitachi SU3500 SEM (A) and observed under vacuum conditions and filament settings of 10-15 kV.

29 Results

Figure caption on page 31.

30

Figure 8: Dissected shoot apical meristems of A. coerulea before (A), during (B-E), and after vernalization (F-I) compared to the nonvernalized control (J). Green arrows signify a vegetative meristem (VM), yellow indicate leaf primordia (LP), white represent an inflorescence meristem (Im), and orange specify an axillary meristem (Am). Red arrows denote lateral floral meristems (LFM) and blue designate the terminal floral meristem (TFM). Meristems were extracted according to the timepoints in table 1. Each letter denotes a collection point during the 8-week experiment. Scale bars are equal to 1.0mm.

31

Figure caption on page 33.

32

Figure 9: Longitudinal sections of A. coerulea before (A), during (B-E), and after vernalization at 6°C (F-I) compared to the nonvernalized control at 24°C (J). Arrows indicate leaf primordia (LP), vegetative meristems (VM), axillary meristems (AM), inflorescence meristems (IB), bracts (Bt), sepals (Sp), stamen primordia (St), terminal buds (TB), and lateral buds (LB) specified by their respective arrows. Slides were stained with the Sharman stain series (Figure 2). Each letter denotes a collection point during the 8-week experiment. Black and red scale bars are set at 100um and 200um, respectively.

33 Figure caption on page 37.

34 Figure caption on page 37.

35 Figure caption on page 37.

36

Figure 10: Plan (capital) and side view (lowercase) orientations of A. coerulea meristems under scanning electron microscopy. Meristems were extracted before (A,a), during (B,b-E,e), and after vernalization at 6°C (F,f-I,i) and compared to nonvernalized control samples at 24°C (J). Sepals of the terminal buds in G-I were removed for analysis of the androecium. Temporal conditions of the meristems are indicated on the lower left corner. The apical meristem is highlighted in blue, emerging leaf primordia in purple, and vegetative structures (stem, leaf, sepal, bracts) in green. The terminal and lateral buds are colored in orange and yellow, respectively. Blue, purple, orange, green, red, yellow, and blue scale bars correspond to their respective measurements. Each letter denotes a collection point during the 8-week experiment.

Figure 11: Comparison of vernalized (V) and nonvernalized (NV) control plants at 4 weeks post-vernalization at 24°C. All plants vernalized at 6°C for 4 weeks produced an inflorescence while nonvernalized control plants kept at 24°C remained vegetative.

37 To investigate the cold-induced floral transition in A. coerulea, morphological studies were conducted at the macro, and microscopic level. Plants were grown to maturity in greenhouse conditions, vernalized at 6°C for 4 weeks, then placed in greenhouse conditions for an additional 4 weeks. The shoot apical meristem (SAM) of several plants were dissected each week of treatment and prepared for SEM (Figure 10

A-J) and histological analysis (Figure 9 A-J). Observations of the dissected shoot apex at pre-vernalization (week 0) revealed a vegetative meristem surrounded by the lower petioles of compound leaves. These whorls of leaves overlapped one another to form a protective barrier around the apical meristem (Figure 3 D; 10 b). Using a stereomicroscope, this rosette of leaves was removed to reveal a vegetative meristem

(Figure 8 A; 9 A).

Under normal, nonvernalized conditions at 24°C, the SAM of A. coerulea produced leaf primordia. These organs formed at the flanks of the meristem underneath the central zone cells. Leaf development could be traced through each rosette with the primordia forming nearest the meristem and more developed forms in the outer whorls.

Budding leaves were observed under SEM and are highlighted in purple (Figure 10 B, b, c). SEM analysis gives a clear view of the developing leaves surrounding the SAM

(highlighted in blue) and the initiation of ternate formation in leaves (highlighted in green) (Figure 10 a, b). In between the leaf epidermal cells are rows of parenchyma cells that make up most of the interior of the leaf tissues. These expand in size as the leaf organ advances in development. Leaf primordia are produced by the SAM during the 1st and 2nd week of vernalization treatment at 6°C (Figure 9 A-C).

38 As seen from the histology results, A. coerulea continues vegetative growth until the 3rd week of vernalization (Figure 9 D). At this length of cold conditions at 6°C, the

SAM itself elongates and develops into an inflorescence meristem (IM), maintaining meristematic activity at the inflorescence apex. Below the IM, axillary meristems (AM) are formed at the flanks immediately below the apex and adjacent to the nearest leaf primordia (Figure 9 D). SEM observations of the shoot apex confirm the formation of lateral AMs (highlighted in yellow) below the IM (Figure 10 D, d). Bract (Bt) formation also begins at 3 weeks of vernalization treatment, with the earliest primordia forming underneath the axillary buds nearest the apex (Figure 9 D). SEM observation confirms the development of bracts subtending the AM (highlighted in light green); however, this is not observed until the 4th week of vernalization (Figure 10 E).

IM identity is maintained throughout the 4th week of vernalization at 6°C. At this stage, the inflorescence begins to branch (Figure 8 D). These new meristems are nested in between leaf and bract primordia that separate them from the main IM. At week 4 of vernalization, branching of these inflorescences was not detected under SEM due to their close association with surrounding leaves that were removed to reveal the apical meristem. After the collection of week 4 vernalized samples, the remaining plants were removed from vernalization conditions in the growth chamber and transferred into greenhouse conditions at 24°C for an additional 4 weeks (Table 1, 4).

After the first week of exposure to warm temperatures (week 1 post- vernalization), observation under the stereomicroscope showed an elongated shoot apex and further development of the terminal and subtending lateral buds (Figure 8 F).

Histological analysis showed a transition in the shoot apex from an IM to a terminal

39 floral meristem (TFM). Lateral AMs remained undifferentiated and continued to undergo cell division at their apices (Figure 8 F). Below the lower whorl of stamen primordia, sepals (Sp) begin to develop at the base of the TFM. Bracts subtending the AMs have elongated significantly at this point of greenhouse treatment. Under SEM, both sepals and bracts (highlighted in green) can be seen subtending the TFM (highlighted in orange) and lateral AMs, respectively (Figure 8 F, f). At the lower end of the TFM, whorls of developing stamens form acropetally towards the apex (Figure 8 F, f).

The second week of greenhouse treatment at 24°C showed the complete formation of the male reproductive structures in the terminal bud. Under histological and

SEM observation, the androecium consisted of several whorls of stamens (Figure 9 G; 8

G). Lateral AMs developed into lateral floral meristems (LFM), which began to develop sepal organs on the flanks below the apex (Figure 8 G). Below the LFMs, secondary

AMs (highlighted in yellow) also began formation flanking the terminal LFM (Figure 9 g). These developing AMs were enclosed by young leaves (highlighted in green) which surrounded the entire inflorescence (Figure 9 g).

At the macroscopic level, developing sepals were observed as they began to enclose the LFMs (red arrows), mimicking the process of sepal growth in the TFM which fully enclosed the apex (blue arrow) (Figure 8 G). Under SEM conditions, sepals (dark green) were clearly seen growing acropetally from their initiation site below the first whorl of stamen primordia (Figure 9 G, g). Sepal organ initiation and development was synchronized across all LFMs at 2 weeks of 24°C conditions, as detected in both stereoscopic and SEM observations (Figure 8 G; 10 G, g).

40 At the third week of greenhouse treatment, the whorls of sepal primordia closest to the apex of the terminal bud separate further into individually lobed organs (Figure 8

H). As seen in histological and SEM microscopy, sepal primordia in the LFMs continued to form acropetally towards the apices (Figure 8 H, 9 H), following the growth pattern of the main terminal bud. Bracts were more developed at the 3rd week of greenhouse conditions in the LFMs and secondary LFMs of the primary inflorescence (Figure 9 H, h). Some LFMs expanded in size and were seen protruding from the subtending leaves.

However, the surfaces remained smooth and free of any stamen primordia, although bract primordia could be seen forming on the flanks of the central zones (Figure 9 H, h).

At the final week of post-vernalization treatment, stereoscopic dissection showed an elongation of the primary inflorescence stem and the surrounding leaves which enclose the developing lateral buds (Figure 8 I). Lateral branches adjacent to the mimicked the pattern of increased height and leaf development in the primary inflorescence. Microscopic analyses showed increased stamen number on the LFMs of the primary inflorescence, nearing the amount of the fully developed terminal bud

(Figure 8 I; 9 I). Similarly, stamen number on the TFMs of the secondary branches also continued to increase (Figure 9 I, i). Sepals in all lateral and terminal floral meristems in all inflorescences grew slightly in size; although they did not completely envelop the developing buds as displayed by the terminal bud on the primary inflorescence (Figure 8

I; 10 I, i).

At the final timepoint of collection (week 9) (Table 1,4), shoot apices of nonvernalized control plants kept at 24°C were dissected for comparing developmental changes with vernalized plants. Observation under the dissecting microscope showed no

41 IM development (Figure 8 A; 9 A; 10 A). Nonvernalized control plants remained in a vegetative state, verified by the production of leaf primordia by the undifferentiated SAM

(Figure 8 J; 9 J; 10 J). After 4 weeks of vernalization treatment at 6°C, plants were moved into greenhouse conditions at 24°C for an additional 4 weeks. By the fourth week of warm temperatures, the primary inflorescence began to emerge from the center of the leaf rosette of all vernalized plants. In comparison, the control plants kept in greenhouse conditions failed to produce flowering stalks (Figure 11).

42 Conclusions

Based on stereoscopic, histological, and SEM analyses of the shoot apex before, during, and after vernalization, mature plants of the A. coerulea variety “Origami Red and White” require vernalization treatment to transition into the reproductive phase. The

SAM remains vegetative and produces leaf primordia until the third to fourth week of vernalization conditions at 6°C (Figure 8 E; 9 D; 10 D). Initial development of lateral

FMs and their subtending bracts occurs from the SAM during vernalization treatment

(Figure 8 E; 9 D, E; 10 D, E).

Inflorescence development increases in complexity in greenhouse conditions (Figure

8 F-I; 9 F-I; 10 F-i). Below the TFM, stamen primordia in the LFMs mimic the acropetal growth pattern of the TFM (Figure 8 F-H; 9 E-H; 10 F-h). Development of the secondary branches initiate during the third to fourth week of vernalization and mimic that of the primary inflorescence (Figure 9 D, E). By the fourth week of greenhouse conditions, buds in the primary inflorescence with compressed branches have produced buds with conspicuous floral organs including stamens, sepals, bracts, and leaves along the elongated inflorescence stalk. In vernalized plants, emergence of the primary inflorescence from the rosette began around 4 weeks post-vernalization in greenhouse conditions (Figure 11).

43 CHAPTER 3

TISSUE CULTURE OF AQUILEGIA COERULEA

Background

In vitro regeneration of Aquilegia coerulea

Model species are important in understanding genetic mechanisms behind evolution and developmental processes in plants. Studying these traits often involve direct genetic manipulation. Establishment of an efficient transgenic protocol is vital for functional studies. Among the many methods of producing transgenic plants, the agrobacterium method of genetic transformation in widely used and standardized for many plants (Kyte, 2013). A plasmid containing the gene of interest is introduced into dedifferentiated callus tissue which then becomes part of the plant genome. Once fully regenerated, the phenotypic effects of transgene can be analyzed. However, current literature lacks tissue culture protocols specific to Aquilegia, restricting further genetic studies.

To develop an efficient tissue culture protocol, empirical optimizations in nutrient, sucrose, and hormonal levels in the media is very important (Kyte, 2013). Since plant species have a diverse range of nutrient and hormone requirements, tissue culture protocols vary for different plant species. Once a tissue culture protocol is established, genetic transformation—a method of introducing novel genes into an organismal genome—can be attempted to obtain stably transformed plants. (Kyte, 2013). This core research tool is instrumental in understanding the function of desired genes in model plant systems like Aquilegia.

44 Objectives & Hypotheses

The objective of this study was to develop efficient protocols for callogenesis shooting and rooting in A. coeruelea. The auxin and cytokinin classes of plant hormones were used in our cultures because of their involvement in growth through cell division and elongation as well as shoot and root organogenesis (Raven, 2012). Through empirical observation and analyses, combinations of auxin and cytokinin plant hormones were tested for their ability to influence induction and growth of undifferentiated callus. Root and shoot culture was subsequently attempted using hormones that showed organogenesis potential in A. coerulea callus.

There are established tissue culture protocols for many model plants, including

Arabidopsis (Sugimoto & Meyerowitz, 2012) and O. sativa (Hoque et al., 2008).

However, few studies have been published on the tissue culture of Aquilegia species.

Work by Fei et al., in 2010 showed that a [0.5/0.6mgL-1] ratio of BAP/2,4-D (6-

Benzylaminopurine/ 2,4-Dichlorophenoxyacetic acid) was optimal for callus induction of

A. oxysepala root explants. Interestingly, plantlets formed in root callus of A. oxysepala

(Fei et al., 2010). Induction of callus tissue will be attempted by experimenting with A. coerulea root, hypocotyl, and cotyledonary tissues. Ratios of synthetic plant hormones

BAP and 2,4-D will be used in combination to assess their effectiveness in callus culture.

Media containing several concentrations of [0.25/0.5/1.0mgL-1] of BAP/2,4-D will be tested. Any combinations found to be conducive to root and shoot formation will be investigated further for enhanced organogenesis in A. coerulea callus tissues.

45 Materials & Methods

In virto seed germination of A. coerulea

To formulate an in vitro regeneration protocol for A. coerulea, sterile plant material was required for experimentation. Seeds of the columbine variety, “Origami Red and White”, were purchased from (swallowtailgardenseeds.com) and used as the main source of explant material for all in vitro experiments. A seed sterilization protocol was developed using solutions of bleach, ethanol, and deionized water (diH2O) in the process described (Figure 13). For the germination media, half strength (17.21gL-1) Murashige and Skoog salt mix with vitamins and sucrose (PhytoTech Labs, cat. no. M5530) was combined with 1000mlL of diH2O in a 1L Erlenmeyer flask (Table 7). After the solution was homogenized with a magnetic stirrer, the pH was adjusted between 5.6-5.8 and high purity agar was added at 6gL-1 (PhytoTech Labs, cat. no. A175). The media was sterilized at 121°C for 30 minutes prior to plating in petri dishes inside the laminar flow hood.

A total of 50 A. coerulea seeds were placed into a 1.5ml microcentrifuge tube.

Using a 70% EtOH solution, the tube was then externally sterilized before being transferred inside the laminar flow hood. Seeds were sterilized by washes in bleach, ethanol, and diH2O as indicated in Table 6. A mini vortexer (Fisher Scientific, cat. no.

02215365) was placed inside the aseptic flow hood and used to fully suspend the seeds in solution at a speed of 1500 RPM. After the final diH2O wash was completed, the seeds were transferred from the microcentrifuge tube onto a glass petri dish containing sterilized filter paper to air-dry. After drying in the laminar flow hood, the seeds were firmly placed in the solidified germination media using sterilized forceps. Plates were wrapped in parafilm and placed under a 16L/8D photoperiod until germination.

46 Table 6: Solutions for sterilization of A. coerulea seeds. A total of 50 seeds were placed into a 1.5ml microcentrifuge tube and with immersed with a 10% bleach solution for 7 minutes followed by 2 minutes of 70% ethanol and diH2O. Sterilized seeds were placed on sterilized filter paper to air-dry before plating on germination media.

Step Solution Vortex time (min)

1 10% bleach 7

2 70% EtOH 2

3 diH2O 2

4 diH2O 2

Figure 12: Sterilization process used for handling all explant and cultured tissues. Forceps and scalpels are first placed in the glass bead sterilizer for 15 seconds (1) then cooled in diH2O (2). Disinfected tools can then be used to cut and transfer explant material (3). Subsequent rinses in 70% ethanol (4) and diH2O (5) ensure sterility is maintained across cultures. The 5-step process is repeated before contact with new plant material.

47 Table 7: A. coerulea germination media: Ingredients and their respective amounts. The half strength MS salt was added to 1000ml of diH2O. The salt solution was adjusted to a pH between 5.6-5.8 using either NaOH or HCL. The agar was then mixed into the solution using a stir bar and magnetic stirrer. The flask was capped and autoclaved at 121° C for 30 minutes.

Germination media components Amount (1L total)

MS salt with vitamins and sucrose 17.2g

Agar 3g

diH2O 1000ml

Figure 13: Diagram of A. coerulea seed sterilization protocol in 6 steps. Seeds are first transferred inside the 1.5ml microcentrifuge tube (1) then washed in 10% bleach (2) and 70% ethanol (3). Sterilized seeds underwent two rinses of diH2O (4,5) before being plated in germination media (6). The blue and orange colored boxes represent the 7 and 2-minute bleach and ethanol solution washes, respectively.

48

Callus culture of A. coerulea

Cotyledons of A. coerulea seedlings germinated in vitro were used as explant material to test the effects of plant hormones BAP and 2,4-D on callus generation. Media in varying concentrations (0.25, 0.5, and 1.0mgL-1) of BAP (6-Benzylaminopurine) and

2,4-D D (2,4-Dichlorophenoxyacetic acid) were formulated using the concentrations in

Table 8. In all BAP/2,4-D combinations, full strength (34.43gL-1) MS basal salt with vitamins and sucrose was used as the source of micro and macro nutrients for the explant

-1 tissues. High purity agar at 6gL was used as the solidifying agent. Deionized H2O

-1 (1000mlL ) was used for media preparation. The MS salt was dissolved with diH2O in a

1L Erlenmeyer flask. After addition of the BAP/2,4-D concentrations (Table 8), the solution was homogenized with a magnetic stir bar and the pH was adjusted between 5.6-

5.8 using NaOH or HCL. The agar was added last and allowed to dissolve into solution.

The flask was then capped and sterilized in an autoclave at 121°C for 30 minutes.

After autoclaving, the flask was placed at room temperature and allowed to cool until it could be held with gloves. Inside the laminar flow hood, the sterile media was poured into plastic petri dishes at 20ml per plate and left to solidify inside the hood.

Using the technique in figure 12, sterilized forceps and scalpels were used to remove the seedlings of A. coerulea from their germination media petri dishes and place them on autoclaved filter paper inside the hood. Using a scalpel, the cotyledons were removed from the hypocotyl, wounded in the leaf midrib area to expose the mesophyll, and placed onto the callus media (Figure 14). Both cotyledons of a single seedling were placed in a single petri dish which served as a biological replicate in all tissue culture experiments.

49 A total of 10 biological replicates per medium treatment were tested. Callus induction and growth in response to hormone ratios was recorded on a weekly basis by photography from a standardized height. Images of cultures were captured at the same focus and depth. From these images, growth of cotyledon area over time was measured and recorded using ImageJ Software (NIH, US). After four weeks, differences in callus area between the hormone treatments was examined.

A B

Figure 14: Dissection of A. coerulea seedlings 2 weeks after germination. In between seedling dissections and subsequent sub-culturing, the scalpel and forceps were cleaned in a glass bead sterilizer for 15 seconds (B) and immersed in solutions of 10% bleach, 70% ethanol, and diH2O.

50 Table 8: Ratios and concentrations of BAP and 2,4-D used in callogenesis of A. coeruela cotyledons. All medias were prepared with 34.43gL-1 MS salt, 6g-1 of agar, 1000ml of diH2O, and respective BAP/2,4-D ratios in a L flask autoclaved at 121°C for 30 minutes. Cultures were placed under a 16L/8D photoperiod at 25°C for 6 weeks. Letters A-I represent the ratios of BAP and 2,4-D.

Medium letter Hormone ratios (mgL-1) BAP (mgL-1) 2,4-D (mgL-1)

A 0.25/0.25 0.25 0.25

B 0.25/0.5 0.25 0.5

C 0.25/1.0 0.25 1.0

D 0.5/0.25 0.5 0.25

E 0.5/0.5 0.5 0.5

F 0.5/1.0 0.5 1.0

G 1.0 1.0 0.25

H 1.0/0.5 1.0 0.5

I 1.0/1.0 1.0 1.0

Figure 15: Protocol for callus culture from A. coerulea seedling cotyledons in 4 steps. After in vitro germination (1), cotyledons of A. coerulea seedlings were dissected and wounded across the leaf rib (2), then firmly placed in the 1/1 BAP/2,4-D callus media (3). Callus generation occurs around 4 weeks after step 3 (4).

51 Based on results of the initial callogenesis study, further experiments were conducted to improve the efficiency of callus growth in wounded cotyledons. The 1/1 ratio of BAP/2,4-D [1/1mgL-1] found to induce undifferentiated callus (Table 13) was doubled to [2/2mgL-1] BAP/2,4-D. These cytokinin and auxin combinations were tested with concentrations of silver nitrate (AgNO3), a compound reported to improve overall growth, shoot organogenesis, and enhanced callus proliferation (Orlikowska, 1997).

Media lacking BAP/2,4-D and AgNO3, as well as medias with only 10, 20, or

-1 -1 50mgL of AgNO3 were used as controls. The dilutions of 10, 20, and 50 mgL

AgNO3were prepared as shown in Table 9. Following a similar method like the initial

BAP/2,4-D callus experiment, both cotyledonary leaves of seedlings germinated in vitro were dissected, wounded, and placed on the media. Both cotyledons of a single seed were placed on a plate of media to serve as a biological replicate. A total of 10 replicates were used for treatments T1-T8 and 6 were used as controls C1-C4 (Table 10). Measurements of callus growth were analyzed using ImageJ software from images of cotyledons taken at weekly timepoints.

Table 9: Concentration of AgNO3 used in callus media. Stock solution was made by dissolving 100mg of AgNO3 in 100ml of diH2O. Solution vials were wrapped in aluminum foil and stored at 25°C.

AgNO3 solution concentration Amount of AgNO3 of Amount of diH2O solvent (mgL-1) stock (ml) (ml)

10 2.5 250

20 5.0 250

50 12.5 250

52 Table 10: Combinations of BAP, 2,4-D, and AgNO3 for callus induction of A. coerulea cotyledons. All media combinations were adjusted to a pH range within 5.60- 5.66 and contained 8.6g of MS salt, 1.5g of agar, and 250ml of diH2O.

-1 -1 -1 Treatment BAP (mgL ) 2,4-D (mgL ) AgNO3 (mgL ) pH

C1 0 0 0 5.61

C2 0 0 10 5.61

C3 0 0 20 5.60

C4 0 0 50 5.61

T1 1.0 1.0 0 5.66

T2 2.0 2.0 0 5.66

T3 1.0 1.0 10 5.60

T4 1.0 1.0 20 5.64

T5 1.0 1.0 50 5.62

T6 2.0 2.0 10 5.64

T7 2.0 2.0 20 5.64

T8 2.0 2.0 50 5.65

53 Organogenesis

By treating A. coerulea cotyledonary leaves with a 1/1 ratio of plant hormones

BAP and 2,4-D, a reliable undifferentiated callus-producing media was established

(Table 13). Formulating a media conducive to the generation of plant shoots and roots from callus was the next phase of experimentation. To this end, calluses generated in medium I ([1/1mgL-1] BAP/2,4-D) (Table 13) were subcultured into medium F and D, which initially showed a high number of root and shoot organogenesis, respectively

(Table 13). Media containing 8.6g of MS salt with vitamins and sucrose, 1.5g of agar, and 250ml of diH2O, was adjusted to a pH range within 5.60-5.65. Rooting and shooting media contained corresponding BAP/2,4-D concentrations of [0.5/1mgL-1] and

[0.5/0.25mgL-1] (Table 11). The media was sterilized at 121°C for 30 minutes using the autoclave. The 20 calluses generated in media I ([1/1mgL-1] BAP/2,4-D) (Table 8) were transplanted into 4 glass vessels divided into rooting and shooting media (Figure 16).

Root and shoot organogenesis was observed and quantified after 9 weeks of culture under a 16L/8D photoperiod at 25°C.

Figure 16: Glass vessels used for root and shoot cultures of A. coerulea callus. Parafilm was wrapped around the openings to prevent contamination from airborne pathogens.

54 Table 11: Combinations of BAP, 2,4-D, used for shoot and root induction in A. coerulea callus tissues. All media was made with 8.6g of MS salt with vitamins and sucrose, 1.5g of agar, 250ml of diH2O, and adjusted to a pH range within 5.60-5.65. Medias were capped and autoclaved at 121°C for 30 minutes.

Hormones Shoot media Root media (mgL-1) BAP 0.5 1.0

2,4-D 0.25 0.5

# of calluses 10 10

Rooting in Liquid Media

Liquid media was used to induce root organogenesis from shooted calluses generated from the shooting media (Table 11). The liquid media was formulated at half- strength MS salt by dissolving 8.6g of the basal salt mix in 250ml of diH2O. After adjustment of the pH between 5.6-5.8, the concentrations of BAP/2,4-D and IAA alone were added (Table 12). The BAP/2,4-D concentration [0.5/1mgL-1] was tested due to its initial high root organogenesis in previous studies (Table 13, medium F). In addition, the synthetic auxin, indole-3-acetic-acid (IAA) was used due to its association with root formation (Leveau & Lindow, 2005). A liquid state was achieved by excluding agar from the media.

A paper bridge method of culture was developed for use with liquid media using a

9cm diameter filter paper (Fisher Scientific, cat. no. 09-795C). The paper was cut in half and folded over on itself several times to form an elongated rectangle. This shape was then bent at both ends and the center to mimic the form of the letter “M” (Figure 18).

Using elongated forceps, the paper bridge was inserted into the tubes of liquid media

55 before autoclave sterilization at 121°C for 30 minutes. After the media was cooled, shooted calluses were introduced into the paper bridge using sterile technique.

Table 12: Treatments of BAP, 2,4-D, and darkness (aluminum foil) used for root induction in A. coerulea shooted callus. All medias were prepared with half strength -1 (17.2g/L ) MS salt, 250ml of diH2O, and their respective concentrations of BAP, 2,4-D, and IAA at a pH of 5.69-5.70. Sterilization took place in an autoclave set at 121°C for 30 minutes. Aluminum foil was wrapped around the entire tube to create a dark environment.

Treatment BAP (mgL-1) 2,4-D (mgL-1) IAA (mgL-1) Darkness pH Tube #

T1 x x 1.0 x 5.70 6

T2 x x 1.0 yes 5.70 6

T3 0.5 1.0 x x 5.69 6

T4 0.5 1.0 x yes 5.69 6

Figure 17: Paper bridge cultures of A. coerulea callus using liquid media. A dark environment was created by wrapping aluminum foil around the tubes. A total of 6 replicates were tested per treatment. Treatment 1 (red cap) = IAA (1mg/L-1); treatment 2 (purple cap) = IAA [1mg/L-1] + dark (7d); treatment 3 (green cap) = [0.5/1mgL-1] BAP/2,4-D; treatment 4 (black cap) = BAP [0.5/1mgL-1] BAP/2,4-D + dark (7d).

56 Shoots derived from [0.5/0.25mgL-1] BAP/2,4-D media (figure 24) were removed inside the flow hood and briefly rinsed in sterile diH2O to remove callus debris. Glass bead-sterilized forceps were used to transfer the shooted calluses and nest them in the furrow of the paper bridge (Figure 18). An additional independent variable of darkness was introduced to both BAP/2,4-D and IAA treatments to test its role in root induction.

Aluminum foil was wrapped around several glass tubes to create a dark environment for the cultures (Figure 17). A total of six replicates were made for each treatment. After 7 days, the foil was removed from the dark-treated tubes. Apart from the foil-covered tubes, all cultures were kept in a photoperiod of 16L/8D for 4 weeks after which root number was counted.

Figure 18: Liquid culture of A. coerulea callus with shoots using a filter paper bridge method. A 9cm diameter filter paper was cut in half, folded rectangularly, and shaped into an “M”. Elongated forceps were used to place the bridge into the tube.

57 Results

In vitro seed germination of A. coerulea

The germination media developed specifically for in vitro seed germination was formulated using half-strength MS concentration at 17.21gL-1 and 6gL-1 of agar to reduce salinity stress for young roots. Together with the sterile technique (Figure 12) and sterilization protocol (Figure 13) established to handle seeds of A. coerulea proved effective. Of the 43 seeds plated, a 97.67% germination rate was reached in 2 weeks under a 16L/8D photoperiod. Seeds showed relatively synchronized germination with radicle emergence taking place at 2 weeks inoculation in germination media. Full cotyledon expansion occurred around 4 weeks in a16L/8D photoperiod at 25°C.

To ensure the efficiency of this protocol, fresh solutions of 10% bleach and 70%

EtOH were made using autoclaved water before each seed cleaning procedure. A total amount of 50 seeds per 1.5ml microcentrifuge tube ensured effective sterilization of all seed surfaces. Seed concentrations of 100 per tube showed increased contamination and reduced germination rates. The finalized protocol for seed sterilization is as shown in

Figure 13.

Explants from in vitro germinated A. coerulea seeds were used for callus induction. To stimulate cell proliferation in the wound sites of seedling tissues, plant hormones auxin 2,4-D (2,4-Dichlorophenoxyacetic acid) and BAP (6-

Benzylaminopurine) purchased from PhytoTechnology Laboratories were used in varying combinations. Sections of root, hypocotyl, and cotyledons dissected from A. coerulea seedlings were wounded using a scalpel and placed on media containing varying ratios of

BAP and 2,4-D (Table 8). A preliminary study indicated that root and hypocotyl tissues

58 were not suitable for culture as they produced minimal callus tissue and succumbed to necrosis within 4 weeks of in vitro culture (Figure 19). Therefore, cotyledons from A. coerulea seedlings were used for callus cultures as they responded best to callus formation and proliferation (Table 13).

Similarly sized cotyledons from seeds that germinated relatively cohesively were harvested. An imaging platform was devised to ensure all images of cultures were taken at equal distance for a consistent resolution. A scale was also included within the frame of each image for proper measurement calibration using the software. However, slight differences in cotyledon size might have affected measurements taken with Image J software.

Figure 19: Dissected hypocotyl of A. coerulea seedling at week 0 (A) and week 4 (B) of callus culture in [0.5/0.5mgL-1] BAP/2,4-D. Although callus was produced in hypocotyl tissues, necrosis was typical at the 4th week of culture. Scale bar represents 1 cm.

59 A

B

Figure caption on page 61.

60 C

Figure 20 A-C: Growth of A. coerulea cotyledon callus tissues (mean ± s.d.) over 6 weeks of culture in BAP/2,4-D media. Growth within treatments at week 0 (A, B) and between treatments at week 6 (C) are compared. Ratios under their respective letters are in mgL-1.

Media A-I varied in their ability to induce callus tissue in cultured cotyledons of

A. coerulea seedlings. Most combinations also promoted the formation of root and shoot organs from callus tissues (Table 13). The concentration of BAP/2,4-D in medium A

[0.25/0.25mgL-1] was doubled to [0.5/0.5mgL-1] in medium E. This resulted in a higher callus area at week 6 of treatment and root production in 40% of cultures. However, when concentrations in medium E were doubled again to [1.0/1.0mgL-1] in medium I, callus remained undifferentiated and slightly decreased in size (Figure 20 B). While maintaining 2,4-D at [0.25mgL-1], a 4-fold increase in BAP concentration in media A, D, and G did not substantially increase callus growth (Figure 20 B). Increasing [BAP] from

61 [0.5mgL-1] in medium D to [1.0mgL-1] in medium G resulted in 40% shoot and 40% root organogenesis, respectively (Table 13).

In a background concentration of [0.25mgL-1] BAP, increased 2,4-D concentrations of [0.25] (medium A), [0.5] (medium B), and [1.0mgL-1] (medium C) promoted minimal root organogenesis (Table 13). There was an increase in callus area as

2,4-D concentration was doubled from [0.5mgL-1] to [1.0mgL-1] in media B and C, respectively (Figure 20 B). In a [0.5mgL-1] BAP background, media D, E, and F had increasing [2,4-D] of 0.25, 0.5, and 1.0mgL-1, respectively (Table 13). Medium D produced shoots in 40% of cultures, but only 70% of cotyledons produced callus.

Medium E achieved 40% rooting which was slightly increased to 44.4% in medium F

(Table 13).

In a constant BAP concentration of [1.0mgL-1], 2,4-D concentration in media G,

H, and I was increased from 0.25, to 5.0 and 1.0mgL-1, respectively. Interestingly, incremental increases in callus size was observed as [2,4-D] concentration was increased

(Figure 20 C). This trend indicated that increasing 2,4-D concentration has a positive effect on callus growth in A. coerulea cotyledons. Callus was produced in all cotyledons in media G, H, and I, with 40% rooting and minimal shooting in medium G, and 30% shooting in medium H. Callus cultures remained undifferentiated in medium I, the only medium to exclusively produce callus tissue in all ratios tested (Table 13).

From the BAP/2,4-D ratios tested, three main combinations were discovered which produced root, shoot, and undifferentiated callus tissues from cotyledons in 6 weeks of culture. Medium F containing a [0.5/1.0mgL-1] ratio of BAP/2,4-D produced callus in 88.90% of cotyledons and root organogenesis in 44.4% of those cultures (Figure

62 21 A, B). Shoot organogenesis was most prevalent in medium D which contained a

[0.5/0.25mgL-1] ratio of BAP/2,4-D (Figure 21 C, D). Shoots were produced in 40% of cultured cotyledons, but callus tissue formed only in 70% of cotyledons (Table 13).

Medium I with a [1.0/1.0mgL-1] ratio of BAP/2,4-D produced undifferentiated callus in all cotyledons (Figure 21 F, E).

Table 13: Ratios of BAP/2,4-D medias A-I and their resulting percentage of induction and organogenesis in A. coerulea cotyledons after 6 weeks of culture. All medias were prepared with full strength (34.43gL-1) MS salt with vitamins and sucrose, -1 6gL of agar, 1000ml of diH2O, and respective BAP/2,4-D ratios in a 1L flask autoclaved at 121°C for 30 minutes. Cultures were placed under a 16L/8D photoperiod at 25°C for 6 weeks. Rooting and shooting percentages across medias are represented by R and S, respectively.

-1 Letter BAP/2,4-D (mgL ) % Callus induction % Organogenesis (n=10) (n=10) A 0.25/0.25 100% 10% R 10%S

B 0.25/0.5 100% 10% R

C 0.25/1.0 100% 20% R

D 0.5/0.25 70% 40% S

E 0.5/0.5 100% 40% R

F 0.5/1.0 88.80% 44.4% R

G 1.0/0.25 100% 40% R 10% S

H 1.0/0.5 100% 30% S

I 1.0/1.0 100% 0%

63 Medium F Medium F

Medium D Medium D

Medium Medium I I Figure 21: Rooting (A, B), shooting (C, D), and callus (F, E) producing media before and after 6 weeks of tissue culture. All organs were formed from callus of A. coerulea seedlings under a 16L/8D photoperiod at 25°C. Scale bars represent 1cm.

64 To reaffirm the potential of organogenesis from A. coerulea cotyledons in media

F, D, and I, further tests were conducted with several additions to the tissue culture media. The [1.0/1.0mgL-1] of BAP/2,4-D produced undifferentiated callus in all cotyledons. The ratio was increased to [2.0/2.0mgL-1] to reduce the time of production and further increase callus size (Table 10). In the initial BAP/2,4-D trial, growth from all callus tissues began to slow, turn brown, and succumb to necrosis past the 6th week of culture; most likely due to the production of phenolic compounds which toxic to the plant tissues (Jones & Saxena, 2013). In the second trial, silver nitrate (AgNO3) was added to the media.

A

Figure caption on page 66.

65

B

C

Figure 22 A-C: Growth of A. coerulea cotyledon callus tissues (mean ± s.d.) over 6 weeks of culture in BAP/2,4-D ± AgNO3 media. Growth within treatments at week 0 (A, B) and between treatments at week 6 (C) are compared. Letters and numbers -1 represent BAP/2,4-D ± AgNO3 media in table 15. Ratios are in mgL .

66 Table 14: Media treatments with their respective ratios of BAP, 2,4-D, and AgNO3. All medias were prepared with full strength (34.43gL-1) MS salt with vitamins and -1 sucrose, 6gL of agar, 1000ml of diH2O, and respective BAP/2,4-D ratios in a 1L flask. Solutions of AgNO3 were added to media after autoclaving at 121°C for 30 minutes. Cultures were placed under a 16L/8D photoperiod at 25°C for 6 weeks.

-1 -1 -1 Treatment BAP (mgL ) 2,4-D (mgL ) AgNO3 (mgL )

C1 0 0 0

C2 0 0 10

C3 0 0 20

C4 0 0 50

T1 1.0 1.0 0

T2 2.0 2.0 0

T3 1.0 1.0 10

T4 1.0 1.0 20

T5 1.0 1.0 50

T6 2.0 2.0 10

T7 2.0 2.0 20

T8 2.0 2.0 50

67 Figure caption on page 71.

68

Figure caption on page 71.

69

Figure caption on page 71.

70

Figure 23 A-X: Cotyledons of A. coerulea before and after 6 weeks of culture in -1 -1 [1/1mgL ] and [2/2mgL ] ratios of BAP, 2,4-D ± AgNO3. All media were prepared -1 -1 with 34.43gL MS salt with vitamins and sucrose, 6gL of agar, 1000ml diH2O, and -1 respective concentrations of 10, 20, and 50mgL AgNO3. Cultures were placed in a 16L/8D photoperiod at 25°C for 6 weeks.

71 After 6 weeks of culture, average callus area was greater in all treatments compared to control media C1-C4 (Figure 22 B). Cotyledons in all control media failed to produce callus tissues and quickly died in vitro within the first week of culture (Figure

23 A-H). The lack of vigor and callus growth in control media show that auxin and cytokine hormones are necessary for inducing cell growth and division. Media containing only MS salt, agar, and AgNO3 was not sufficient to sustain the growth and vitality of excised A. coerulea cotyledons (Figure 23 A-H).

Medium T1, which contained the [1.0/1.0] ratio of BAP/2,4-D, produced the highest amount of callus compared to all other media with an average size of 70mm2

(Figure 22 B). The lack of AgNO3 in the medium may have added to the considerable difference in growth between week 0 and week 6 of treatment. The callus produced in medium T1 remained undifferentiated, similar to the callus in the BAP/2,4-D study

(Figure 21 E). The error bars in medium T1 represent a large range of standard deviation which may be due to outliers of a higher or lower callus area.

Pairwise comparisons indicated that callus area within treatments T1-T8 were greater after 6 weeks of culture compared to controls C1-C4 which did not see a large increase in callus area (Figure 22 C). Excluding the control media, the greatest and lowest callus area produced after the 6 week of treatment was produced in medium T1 and T3, respectively (Figure 22 C).

Doubling the BAP/2,4-D to [2.0/2.0mgL-1] in media T2 did not result in callus area that surpassed that produced by media T1 [1.0/1.0mgL-1]. After 6 weeks of culture, callus area decreased in media T2, suggesting there is a negative effect of high auxin and cytokinin concentrations and callus proliferation (Figure 22 C). Interestingly,

72 [2.0/2.0mgL-1] BAP/2,4-D media T6, T7, and T8 saw incremental increases in callus area

-1 with the addition of 10, 20, and 50mgL AgNO3 respectively (Figure 22 B). A similar

-1 pattern of increasing callus growth and AgNO3 concentration was seen in 1.0/1.0mgL

[BAP/2,4-D] T3 and T4 media, although callus area slightly decreased in media T5.

-1 The introduction of AgNO3 into the tissue culture media at 10, 20, or 50mgL did not have an effect on either callus growth (Figure 23 A-X). As seen in the BAP/2,4-D experiments without AgNO3, calluses began to brown and succumb to necrosis around the 6th week of culture (Figure 23 J, L). At the same timepoint, similar browning was seen in all callus tissues regardless of AgNO3 concentration (Figure 23 N, P, R, T, V, X).

Media with AgNO3 saw a change in color, appearing a darker shade of yellow-brown with increasing concentrations in both [1.0/1.0mgL-1] (Figure 23 N, P, R) and

[2.0/2.0mgL-1] BAP/2,4-D ratios (Figure 23 T, V, X). Tissue browning in all media suggested that nutrients were quickly up-taken and utilized by the growing callus tissues.

Subculturing, the periodic transfer of in vitro plant tissues into fresh media, may be a useful technique (Kyte, 2013).

Based on A. coerulea callus observations, the health of callus tissues peaked near the fourth week of culture, after which browning and necrosis began to appear (Figure 23

N, P, R, T, V, X). In future in vitro experiments, healthy callus should be dissected at the fourth week of culture, and immediately transferred into a fresh plate of media. If done under sterile conditions, this propagation method can ensure a steady supply of A. coerulea callus without the time and labor required for in vitro seed germination to obtain cotyledon explants. Both BAP/2,4-D ± AgNO3 studies showed potential for undifferentiated callus with [1.0/1.0mgL-1] BAP/2,4-D in cotyledons.

73 Shoot and root induction of A. coerulea callus

By treating A. coerulea cotyledons with a [1.0/1.0mgL-1] ratio of plant hormones

BAP and 2,4-D, a reliable callus medium was established. Formulating a medium conducive to the generation of shoots and roots from callus was the next phase of experimentation. To this end, media with the highest rooting and shooting frequency were determined using callus generated by using cotyledons as explants. In place of narrow petri dishes, glass vessels were used to accommodate the vertical growth of roots and shoots (Figure 16).

In the initial callogenesis experiment, cultures of medium F [1.0/0.25mgL-1] and

D [0.5/0.25mgL-1] achieved a 44.4% root and 40% shoot organogenesis respectively after

6 weeks of culture in a 16L/8D photoperiod at 25°C (Table 13). To improve root and shoot induction, cultures were placed in the same conditions for a longer 9 week period to allow for root and shoot organogenesis and elongation. A total of 10 calluses per media treatment were embedded in four vessels divided into two shooting and two rooting media treatments (Figure 16). Due to the nature of the glass vessel, length measurements were not able to be performed. Instead, the number of organs produced per vessel were quantified after the conclusion of the 9 week time period.

At the end of 9 weeks, callus tissues in the vessels containing shooting media of

[0.5/0.25mgL-1] BAP/2,4-D produced a total of 50 shoots (Figure 24). Interestingly, 22 shoots were also formed in the rooting media (Table 15). Despite use of an aseptic technique inside a sterile flow hood, some cultures became contaminated and were quickly removed from the vessels. At the end of the time period, 6 shooting and 8 calluses remained uncompromised (Table 15). Average number or shoot organs produced

74 per callus was 8.33 in shoot media and 2.75 in root media. Interestingly, no roots were produced in the rooting media which saw a 44.4% frequency of root organogenesis in the initial BAP/2,4-D study (Table 13).

The shooting potential of 0.5/0.25mgL-1 [BAP/2,4-D] was confirmed from the results of this study. After a 9 week period, shoots were around 2-3 inches in length. In future studies, transfer of the entire callus into a rooting media would be the next step toward complete plant regeneration. Interestingly, rooting failed to occur in the media which produced roots in the initial study. Perhaps increasing bioreplicates, subculturing into other identified rooting media combinations of BAP/2,4-D (media E [0.5/0.5mgL-1]; media H [1.0/0.25mgL-1]) or use of other root promoting hormones like IAA would induce rooting more successfully.

Table 15: Ratios of BAP/2,4-D in callus medias and their resulting percentage of shoot and root induction in A. coerulea cotyledons after 9 weeks of culture. *Number of disease-free calluses remaining out of the initial 10 calluses.

-1) Hormones (mgL Shoot media Root media

BAP 0.5 1.0

2,4-D 0.25 0.25

# of calluses 6* 8*

Total shoot # 50 22

Total root # 0 0

Average organ # per callus 8.33 2.75

75

Figure 24: Shoot organogenesis from A. coerulea callus tissues in [0.5/0.25mgL-1] BAP/2,4-D after 9 weeks of culture in a 16L/8D photoperiod at 25°C. Number of shoots produced were quantified visually after removal of compromised callus tissues

Liquid root culture of A. coerulea shoots

Poor rooting in solid media with a 44.4% root induction frequency in the

BAP/2,4-D study (Table 13) led to testing the suitability of liquid media for rooting. A filter paper bridge method of culture (Figure 18) was used to allow contact of the liquid media and the shoot cultures derived from the previous organogenesis study (Figure 24).

The rooting ratio of [0.5/1.0mgL-1] BAP/2,4-D was tested again, this time with the exclusion of agar and the addition of dark treatment for 7 days. Tule et al. (2005) reported that 2,4-D can be used in combination with BAP for induction of rooting in explants with multiple shoots. Indole-3-acetic-acid (IAA) was also tested due to its involvement in light responses and high sensitivity by root tissues (Leveau & Lindow, 2005). An IAA concentration of [1.0mgL-1] was used in liquid media in both light and 7 days of dark treatment to test for effects on root organogenesis from shooted callus tissues. Due to minimal shooted callus material, hormone free controls were not used in this preliminary study.

76

Figure 25: Adventitious root organogenesis from A. coerulea shooted calluses in [1.0mgL-1] IAA + 7 days of dark treatment. Cultures were placed under a 16L/8D photoperiod at 25°C, except for cultures treated with 7 days of darkness. Number of roots produced from calluses was checked at the fourth week of culture.

Table 16: Number of roots produced from A. coerulea shooted calluses in BAP/2,4-D and IAA liquid media ± 7 days of darkness. Total roots produced from calluses were observed using a stereomicroscope and quantified.

Treatment BAP (mgL-1) 2,4-D (mgL-1) IAA (mgL-1) Darkness Root #

T1 x x 1.0 x 4

T2 x x 1.0 yes 24

T3 0.5 1.0 x x 0

T4 0.5 1.0 x yes 0

77 Cultures were observed for root organogenesis after four weeks of culture in a

16L/8D photoperiod at 25°C. Liquid media containing [1.0mgL-1] IAA + 7 day dark treatment produced more roots than IAA in light. Concentrations of [0.5/1.0mgL-1] BAP and 2,4-D failed to produce roots in either light or dark treatments (Table 16). Based on observational analysis of the paper bridge cultures under a stereoscope, adventitious root formation was enhanced under a liquid IAA media and a dark treatment of 7 days (Figure

25). Interestingly, the media which initiated rooting from A. coerulea cotyledon callus in the initial BAP/2,4-D study (Table 13) failed to induce any root organogenesis in a liquid form, even in conjunction with the 7 day dark treatment (Table 16).

Although initially healthy, the residual callus at the base of shoots turned brown prior to transfer into tubes containing a paper bridge in liquid media. The moistened paper bridge indicated that nutrient media was accessible to the callus cells. Our study attempted rooting in calluses containing only 1 or 2 shoots in limited hormone combinations (Figure 18). Darkness seems to have a role in the induction of roots from callus tissues. Further studies will investigate how longer exposures to darkness under additional concentrations of IAA affect root organogenesis in A. coerulea callus.

78 Conclusions

Several experiments were conducted to formulate an in vitro plant regeneration protocol specifically for A. coerulea. To prevent contamination, seeds of the columbine variety “Origami Red and White” were sterilized and plated on germination media.

Synchronous and uniform seed germination was achieved in 2 weeks. After producing sterile plant tissue for in vitro experimentation, ratios of plant hormones 2,4-D (auxin) and BAP (cytokinin) were formulated into various media for callus production. Seedling root, hypocotyl, and cotyledon tissues were firmly placed on media and observed for callus induction and growth. Of these, only cotyledons showed potential to produce callus under controlled conditions. Roots and hypocotyls quickly experienced necrosis in vitro.

Subsequent experiments were conducted with BAP/2,4-D ratios with and without

AgNO3 to reduce toxic phenolic compounds detrimental to callus health. No significant effects on either callus proliferation or phenolic browning were observed after 6 weeks.

Based on these data, a [1.0/1.0mgL-1] ratio of BAP/2,4-D was found to produce reliable, undifferentiated callus tissue. From callus, shooting and rooting were attempted using media formulated with hormone ratios which correlated with organogenesis. Of these, a ratio of [0.5/0.25mgL-1] BAP/2,4-D resulted in prolific shooting rate after a 9 week period. Liquid media was used to induce rooting from shooted callus nested in a paper bridge. A concentration of [1.0mgL-1] IAA in conjunction with 7 days of dark treatment was found to induce adventitious root formation in shooted calluses. These experiments outline the formation of an effective in vitro regeneration protocol for A. coerulea.

Further work aims to refine media for increased organogenesis and complete regeneration of A. coerulea.

79 Few studies have previously explored in vitro regeneration of Aquilegia species and to our knowledge, none have published tissue culture methods specific to A. coerulea. In 2016, Ahmad et al. attempted to microprogate, A. nivalis, member of the

Aquilegia genus native to the Himalaya mountain range known for its use in traditional medicines. Combinations of NAA (1-Naphthaleneacetic acid), 2,4-D, kinetin, and BAP in MS media, were tested with leaf (petiole, middle, and tip regions), shoot tips, roots, and petioles of mature A. nirvalis plants. Callus formation was induced from petiole tissues in 29 days on MS medium containing [1.5mgL-1] NAA and [1mgL-1] kinetin.

Callus was responsive to subcultures in MS medium with 2,4-D [2mgL-1] and kinetin

[2mgL-1] (Ahmad et al., 2016).

Another micropropagation study was conducted by Fei et al. in the species A. oxysepala, a perennial herb rich in alkaloids and flavonoids used in Chinese medicine.

High seed sterility was attained after a 75% alcohol rinse (30 seconds) followed by 5 water rinses, and a 0.2% mercuric chloride rinse (2 min). Following sterilization, germination rate reached 100% on MS medium containing 3% sugar and 0.6% agar (Fei et al., 2010). Although stem and leaf segments were also tested, callus proliferation was highest (92%) in root cultures containing of MS, 2,4-D [0.6mgL-1] and BAP [0.5mgL-1].

Plantlets were formed from the root callus, showing potential for regeneration whole plant (Fei et al., 2010). No specific media pH was indicated in the conclusions by Fei and colleagues. However, based on soil guidelines from the Syngenta culture guide, we adjusted the media pH between 5.6-5.8, which was conducive to the growth of A. coerulea tissues in vitro.

80 In our in vitro culture of A. coerulea, explant tissues from aseptically germinated seed were chosen for ensured sterility. Using our sterilization method (Figure 13), seed germination reached 97.67% after 2 weeks of culture in germination media (Table 7).

Callus induction was best achieved using cotyledons as explants in treatments of phytohormones 2,4-D (auxin) and BAP (cytokinin).

Phytohormones used across Aquilegia in vitro studies were primarily synthetic auxins and cytokines. In our studies, combinations of 2,4-D and BAP induced callus proliferation in all ratios tested, with certain combinations resulting either root or shoot organogenesis or undifferentiated callus (Table 13). In vitro studies of Aquilegia species from Ahman et al. and Fei et al. did not report root or shoot organogenesis from their callus tissues. We found that MS medium containing a balanced combination of [1/1mgL-

1] BAP/2,4-D resulted in undifferentiated callus production in cotyledons after 4 weeks of culture, a timeframe comparable to petiole callus in A. nivalis shown by Ahmad et al.

(2016). Furthermore, shoot organogenesis was induced using [0.5/0.25mgL-1] BAP/2,4-

D). Root organogenesis was high in liquid media with MS and [1mgL-1] IAA in conjunction with a 7-day dark treatment (Table 16, Figure 25).

To our knowledge, our tissue culture of callus, root, and shoot organs represent the first of such studies specific to A. coerulea. Previous authors have successfully induced callus in both A. nivalis and A. oxysepala, although explant tissues varied from petiole to roots, respectively (Ahmad et al., 2016; Fei et al., 2010). Callogenesis and organogenesis response is dependent on explant tissue and media composition.

81 REFERENCES

Ahmad, M., Kaloo, Z. A., Ganai, B., Ganaie, H. A., & Padder, B. M. (2016). Callus Induction Studies in Aquilegia nivalis Flac Ex Jackson: An Endangered Medicinal Plant of Kashmir Himalaya: A Perspective. Journal of Plant Sciences,11(1), 31- 37. doi:10.3923/jps.2016.31.37

Amasino, R. (2004). Vernalization, Competence, and the Epigenetic Memory of Winter. The Plant Cell Online,16(10), 2553-2559. doi:10.1105/tpc.104.161070

Amasino, R. (2005). Vernalization and Flowering Time. Current Opinion in Biotechnology 16: 154–58.

Amasino, R. (2010). Seasonal and developmental timing of flowering. The Plant Journal, (61), 1001–1013. https://doi.org/10.1111/j.1365-313X.2010.04148.x

Amasino, R. M., & Michaels, S. D. (2010). The Timing of Flowering. Plant Physiology,154(2), 516-520. doi:10.1104/pp.110.161653

Alexandre, C. M., & Hennig, L. (2008). FLC or not FLC: the other side of vernalization. Journal of Experimental Botany, 59(6), 1127–1135.

Ballerini, E., & Kramer, E. (2011). Environmental and Molecular Analysis of the Floral Transition in the Lower Eudicot Aquilegia Formosa. EvoDevo, 2(4):20. https://doi.org/10.1186/2041-9139-2-4

Berendse, F., & Scheffer, M. (2009). The angiosperm radiation revisited, an ecological explanation for Darwin’s “abominable mystery.” Ecology Letters, 12(9), 865– 872. https://doi.org/10.1111/j.1461-0248.2009.01342.x

Bi, M., Lijuan, W., Annaliese, M., Zeshan, A., & Meili, X. (2013). Characterization and comparison of key genes involved with flowering time regulation from Arabidopsis thaliana, Oryza sativa and Zea mays. African Journal of Biotechnology, 12(4), 353–363. https://doi.org/10.5897/AJB12.1778

Blázquez, M. A., Green, R., Nilsson, O., Sussman, M. R., & Weigel, D. (1998). Gibberellins Promote Flowering of Arabidopsis by Activating the LEAFY Promoter. The Plant Cell, 10(5), 791–800. https://doi.org/10.1105/tpc.10.5.791\

Brown P, Klein P, Bortiri E, Acharya C, Rooney W, Kresovich S. (2006). Inheritance of inflorescence architecture in sorghum. Theoretical and Applied Genetics. 113:931–942.

Caicedo, A. et al. (2004). Epistatic Interaction between Arabidopsis FRI and FLC Flowering Time Genes Generates a Latitudinal Cline in a Life History Trait. PNAS 101(44): 15670–75.

82 Caspari, E. W., & Marshak, R. E. (1965). The Rise and Fall of Lysenko. Science,149(3681), 275-278. doi:10.1126/science.149.3681.275

Chouard, P. (1960). Vernalization and its Relations to Dormancy. Annu Rev Plant Physiol 11:191-238.

Deng, W. et al. (2011). FLOWERING LOCUS C (FLC) Regulates Development Pathways throughout the Life Cycle of Arabidopsis. PNAS 108(16): 6680–85.

Fei, W., Si-jia, J., Zhong-cai, L., & Xiao-dong, L. (2010). Study on Explants Sterilization and Callus Induction of Aquilegia xysepala. Agricultural Science & Technology,11(7), 25-28.

Fernando, A., and Coupland, G. 2012. “The Genetic Basis of Flowering Responses to Seasonal Cues.” Nature Reviews Genetics 13(9): 627–39.

Fornara, F., de Montaigu, A., & Coupland, G. (2010). SnapShot: Control of Flowering in Arabidopsis. Cell, 141(3), 550–550.e2. https://doi.org/10.1016/j.cell.2010.04.024

Franks, S. J., Sim, S., & Weis, A. E. (2007). Rapid evolution of flowering time by an annual plant in response to a climate fluctuation. Proceedings of the National Academy of Sciences, 104(4), 1278–1282. https://doi.org/10.1073/pnas.0608379104

Garner, W. W., & Allard, H. A. (1920). Effect of the relative length of day and night and other factors of the environment on growth and reproduction in plants. Monthly Weather Review, 48(7), 415–415. https://doi.org/10.1175/15200493(1920)48

Gassner, G. (1918). Beitraege Zur Physiologischen Charakteristik Sommer-und Winterannueller Gewaechse in Besondere der Getreidepflanzen. Zeitschrift für Botanik 10: 27-476.

Glover, B. (2014). Understanding Flowers and Flowering (2nd ed.). Oxford University Press.

Gould, B. Kramer, E. (2007). Virus-induced gene silencing as a tool for functional analyses in the emerging model plant Aquilegia (columbine, Ranunculaceae). Plant Methods 3:1-12

Hoque, M., Ali, M., & Karim, N. (2008). Embryogenic Callus Induction and Regeneration of Elite Bangladeshi Indica Rice Cultivars. Plant Tissue Culture and Biotechnology,17(1). doi:10.3329/ptcb.v17i1.1122

Huang, J., Pray, C., & Rozelle, S. (2002). Enhancing the crops to feed the poor. Nature, 418(6898), 678.

83

Jones, A. M., & Saxena, P. K. (2013). Inhibition of Phenylpropanoid Biosynthesis in Artemisia annua L.: A Novel Approach to Reduce Oxidative Browning in Plant Tissue Culture. PLoS ONE, 8(10). doi:10.1371/journal.pone.0076802

Jordan, B. R. (2006). The Molecular Biology and Biotechnology of Flowering. Wallingford, UK; Cambridge, MA: CABI Pub.

Jung, C., Wong, C., Singh, M. B., & Bhalla, P. L. (2012). Comparative Genomic Analysis of Soybean Flowering Genes. PLoS ONE, 7(6), 1–16.

Jung, C., & Müller, A. E. (2009). Flowering time control and applications in plant breeding. Trends in Plant Science, 14(10), 563–573. https://doi.org/10.1016/j.tplants.2009.07.005

Kawamura, K., & et, al. (2011). Quantitative trait loci for flowering time and inflorescence architecture in rose. Theoretical & Applied Genetics. 122: 661–675.

Klejnot, John, and Chentao Lin. (2004). A CONSTANS Experience Brought to Light. Science 303(5660): 965–66.

Koornneef, M., Hanhart, C.J., & van der Veen, J.H. (1991). A Genetic and Physiological Analysis of Late Flowering Mutants in Arabidopsis thaliana. Mol. Gen. Genet. 229: 57–66.

Knott, J. E. (1934). Effect of a Localized Photoperiod on Spinach. Proc. Amer. Soc.Hort. Sci. 31:152-154.

Kramer, E., Jaramillo, A., & Di Stilio, V. (2004). Patterns of Gene Duplication and Functional Evolution During the Diversification of the AGAMOUS Subfamily of MADS Box Genes in Angiosperms. Genetics, 166(2), 1011–1023. https://doi.org/https://doi.org/10.1534/genetics.166.2.1011

Kramer, E. M. (2009). Aquilegia: A New Model for Plant Development, Ecology, and Evolution. Annual Review of Plant Biology, 60, 261–277. https://doi.org/10.1146/annurev.arplant.043008.092051

Kwiatkowska, D. (2008). Flowering and apical meristem growth dynamics. Journal of Experimental Botany, 59(2), 187– 201.https://doi.org/https://doi.org/10.1093/jxb/erm290

Kyte, L. (2013). Plants from test tubes: An introduction to micropropagation. Portland, OR: Timber Press.

Lang, A. (1965). Physiology of flower initiation. In Encyclopedia of Plant Physiology, W. Ruhland, ed (Berlin: Springer-Verlag), pp. 1371–1536.

84

Lang, A., Chailakhyan, M. K., & Frolova, I. A. (1977). Promotion and inhibition of flower formation in a dayneutral plant in grafts with a short-day plant and a long- day plant. Proc. Natl. Acad. Sci. 6: 2412–2416.

Langridge, J. (1957). Effect of Day-length and Gibberellic Acid on the Flowering of Arabidopsis. Nature, 180(4575), 36–37. https://doi.org/10.1038/180036a0

Leveau, J. H., & Lindow, S. E. (2005). Utilization of the Plant Hormone Indole-3-Acetic Acid for Growth by Pseudomonas putida Strain 1290. Applied and Environmental Microbiology,71(5), 2365-2371. doi:10.1128/aem.71.5.2365-2371.2005

Lin, C., & Lin. (2000). Photoreceptors and regulation of flowering time. Plant Physiology, 123(1), 39.

Lysenko, T. D. (1928). Effect of the Thermal Factor on the Duration of the Developmental Phases of Plants. Experiments with Cereals and Cotton. Im. Tov. Ordzhonikidze (Baku) 3.

Machida, Y., Fukaki, H., & Araki, T. (2013). Plant Meristems and Organogenesis: The New Era of Plant Developmental Research. Plant and Cell Physiology,54(3), 295- 301. doi:10.1093/pcp/pct034

Mandoli, D. F., & Olmstead, R. (2000). The Importance of Emerging Model Systems in Plant Biology. Journal of Plant Growth Regulation, 19, 249–252. https://doi.org/10.1007/s003440000038

Markovskaya, E.F., & Sysoeva, M.I. (2011). Evolution of Plant Photoperiodism. Acta Horticulturae. 907:189-192

Michaels, S.D., Amasino, R. FLOWERING LOCUS C encodes a novel MADS domain protein that acts as a repressor of flowering. Plant Cell, 11 (1999), pp. 949–956

Napp-Zinn, K. (1987). Vernalization: Environmental and genetic regulation. In Manipulation of Flowering, J.G. Atherton, ed (London: Butterworths), pp. 123– 132.

Orlikowska, T. (1997). Silver Nitrate Inhibits Bacterial Growth in Plant Tissue Cultures. Agricell Report,29(4), 96-100.

Pabón-Mora, N., Sharma, B., Holappa, L. D., Kramer, E. M. and Litt, A. (2013). The Aquilegia FRUITFULL-like genes play key roles in leaf morphogenesis and inflorescence development. Plant Journal 74: 197–212.

85 Putterill, J., Robson, F., Lee, K., Simon, R., & Coupland, G. (1995). The constans gene of arabidopsis promotes flowering and encodes a protein showing similarities to zinc finger transcription factors. Cell, 80(6), 847.

Raven, P. H., Eichorn, S. E., & Evert, R. F. (2012). Raven Biology of Plants: 8th ed. New York: W.H. FREEMAN & CO.

Ream, T. S., Woods, D. P., & Amasino, R. M. (2012). The Molecular Basis of Vernalization in Different Plant Groups. Cold Spring Harbor Symposia on Quantitative Biology,77(0), 105-115. doi:10.1101/sqb.2013.77.014449

Rolland, F., Baena-Gonzalez, E., & Sheen, J. (2006). Sugar Sensing and Signaling in Plants: Conserved and Novel Mechanisms. Annual Review of Plant Biology, 57, 676–699. https://doi.org/10.1146/annurev.arplant.57.032905.105441

Sharma, B., Guo, C., Kong, H., & Kramer, E. M. (2011). Petal-specific subfunctionalization of an APETALA3 paralog in the Ranunculales and its implications for petal evolution. New Phytologist,191(3), 870-883. doi:10.1111/j.1469-8137.2011.03744.x

Samach, A., Onouchi, H., Gold, S., Ditta, G., Schwarz-Sommer, Z. , et al. (2000). Distinct roles of constans target genes in reproductive development of arabidopsis. Science, 288(5471), 1613-1616.

Shedron, K. G., & Weiler, T. C. (1982). Regulation of growth and flowering in Aquilegia X hybrida Sims. J. Amer. Soc. Hort. Sci, 107, 878-882.

Simpson, G. G., & Dean, C. (2002). Arabidopsis, the Rosetta Stone of Flowering Time? Science, 296(5566), 285–289.

Simpson, G. G. (2004). The autonomous pathway: epigenetic and post-transcriptional gene regulation in the control of Arabidopsis flowering time. Current Opinion in Plant Biology, 7(5), 570–574. https://doi.org/10.1016/j.pbi.2004.07.002

Son, H., Park, B., Song, J., & Seo, H. (2014). FLC-Mediated Flowering Repression Is Positively Regulated by Sumoylation. Journal of Experimental Botany 65(1): 339–51.

Souer, E., van der Krol, A.R., Kloos, D., Spelt, C., Bliek, M., Mol, J., and Koes, R. (1998). Genetic control of branching pattern and floral identity during petunia inflorescence development. Development 125: 733–742.

Strange, A., Li, P., & Anderson, A. (2011). Major-Effect Alleles at Relatively Few Loci Underlie Distinct Vernalization and Flowering Variation in Arabidopsis Accessions. PLoS ONE, 6: 1–11. https://doi.org/10.1371/journal.pone.0019949

86 Sugimoto, K., & Meyerowitz, E. M. (2012). Regeneration in Arabidopsis Tissue Culture. Methods in Molecular Biology Plant Organogenesis,265-275. doi:10.1007/978-1-62703-221-6_18

Sung, S., & Amasino, R. (2004). Vernalization and epigenetics: how plants remember winter. Current Opinion in Plant Biology, 7, 4–10 https://doi.org/10.1016/j.pbi.2003.11.010

The plant list (2013). The Angiosperms (Flowering plants) [Data file]. Retrieved from: http://www.theplantlist.org/

Tule, D., Ghorade, R. B., Mehatre, S., Pawar, B. V., & Shinde, E. (2005). Rapid multiplication of tumeric by micropropagation. Ann. Plant Physiol.,19, 35-37.

Upadyayula N, da Silva H, Bohn M, Rocheford T. (2006). Genetic and QTL analysis of maize tassel and ear inflorescence architecture. Theoretical and Applied Genetics. 112:592–606.

Valverde, F. et al. (2004). Photoreceptor Regulation of CONSTANS Protein in Photoperiodic Flowering. Science 303(5660): 1003.

Vijayraghavan, U., Prasad, K., & Meyerowitz, E. (2005). Specification and maintenance of the floral meristem: interactions between positively-acting promoters of flowering and negative regulators. Current Science, 89(11), 1835–1843.

Wellensiek, S. (1973). Effects of vernalization and gibberellic acid on flower bud formation in different genotypes of pea under different photoperiods. Scientia Horticulturae,1(2), 177-192. doi:10.1016/0304-4238(73)90029-0

White, J. W., Chen, H., Zhang, X., Beattie, D. J., & Grossman, H. (1990). Floral Initiation and Development in Aquilegia. HortScience,25(3), 294-296.

Wilson, R. N., Heckman, J. W., & Somerville, C. R. (1992). Gibberellin Is Required for Flowering in Arabidopsis thaliana under Short Days. Plant Physiology, 100(1), 403–408. https://doi.org/10.1104/pp.100.1.403

Yamaguchi, N., Winter, C. M., Wu, M.-F., Kanno, Y., Yamaguchi, A., Seo, M., & Wagner, D. (2014). Gibberellin Acts Positively Then Negatively to Control Onset of Flower Formation in Arabidopsis. Science, 344(6184), 638. https://doi.org/10.1126/science.1250498

Yan, L. (2004). The Wheat VRN2 Gene Is a Flowering Repressor Down-Regulated by Vernalization. Science,303(5664), 1640-1644. doi:10.1126/science.1094305

87 Xiang, L., Li, X., Qin, D., Guo, F., Wu, C., Miao, L., & Sun, C. (2012). Functional analysis of FLOWERING LOCUS T orthologs from spring orchid (Cymbidium goeringii Rchb. f.) that regulates the vegetative to reproductive transition. Plant Physiology and Biochemistry, 58, 98–105. https://doi.org/10.1016/j.plaphy.2012.06.011

88