METABOLIC ENGINEERING OF PLANTS BY MANIPULATING POLYAMINE TRANSPORT AND BIOSYNTHESIS
Sheaza Ahmed
A Dissertation
Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
December 2017
Committee:
Paul Morris, Advisor
Andrew T Torelli Graduate Faculty Representative
Vipa Phuntumart
Scott Rogers
George Bullerjahn
ii ABSTRACT
Paul Morris, Advisor
Transport is an essential component of the regulation of polyamines, but to date only one family of Polyamine Uptake Transporters (PUTs) have been characterized in plants, and their impact on polyamine regulation has not been defined. Here we show that knockout mutants of put5 in A. thaliana, promote early flowering and result in plants with smaller leaves, thinner stems, and fewer flowers. In contrast, heterologous expression of the rice gene OsPUT1 in A. thaliana using the Put5 promoter at 22°C produced plants with larger leaves, a two-week delay of flowering and more flowers and siliques. Similar effect on leaf size, flowering time and number of siliques also were observed in transgenic plants with constitutive expression of
OsPUT1 or OsPUT3. The delay of flowering was associated with significantly higher levels of spermidine and spermidine conjugates in the leaves prior to flowering. These experiments outline the first genetic evidence for the control of flowering by polyamines. How polyamine levels control the timing of flowering at a molecular level is not yet known, but this delay of flowering has been demonstrated to be upstream of the stimulation of flowering by the gibberellin and temperature sensitive response pathways.
It has been assumed that there exists a single cytosolic pathway for the synthesis of putrescine in A. thaliana. Here we show that A. thaliana and Glycine max, have a chloroplast- localized putrescine biosynthetic pathway. This pathway comprises of arginine decarboxylase
iii and an agmatinase to synthesize putrescine from arginine. Analysis of expression data suggests that it is the major route of putrescine synthesis in response to stress signals.
Since compartmentation of polyamines has been demonstrated to play an essential role in polyamine homeostasis, the identification of other types of polyamine transporters is a critical knowledge gap. We show here that PDR11 is an important long-distance transporter of polyamines in plants and that OCT5 functions as a vacuolar transporter for polyamines. Taken , these findings will accelerate interest in manipulating polyamine metabolism to generate more stress responsive crop plants.
iv
Dedicated to my wonderful and loving family.
v ACKNOWLEDGMENTS
The last five years has been a period of intense learning for me, not only in the scientific filed, but on a personal level as well. I would like to thank all the people who have supported and helped me throughout this period.
Firstly, I would like to express my sincere gratitude to my advisor Dr. Paul Morris for the continuous support and guidance during my Ph.D., for his patience, motivation, and immense knowledge. His guidance has helped me during the time of research and in writing of this dissertation. I could not have imagined having a better advisor and mentor than him.
Besides my advisor, I would like to thank the rest of my dissertation committee members:
Dr. Vipa Phuntumart, Dr. Scott Rogers, Dr. George Bullarjahn and Dr. Andrew Torelli for their insightful comments and encouragement, but also for the hard question which incented me to widen my research from various perspectives. I would also like to thank the faculty and staff members of the biology department for being very helpful and supportive.
I thank my fellow lab mates: Dr. Jigar Patel, Dr. Lingxiao Ge, Menaka Ariyaratne,
Chandra Sarkar, Nilanjana Chakrabarti and all the amazing undergraduate students who worked with me on various projects. I am extremely grateful for the stimulating discussions, endless help and support, and for all the fun we have had in the last five years.
Last but not the least, I would like to thank my family: my parents and my brothers for supporting me spiritually throughout my Ph.D. and my life in general.
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TABLE OF CONTENTS
Page
CHAPTER I. INTRODUCTION ...... ……………………… 1
1.1. What Are Polyamines? ...... …………………………………………. 1
1.1.1. Polyamine Biosynthesis ...... 2
1.1.2. Polyamine Catabolism ...... 4
1.1.3. Polyamine Transport ...... 6
1.2. Polyamine and Development ...... 8
1.2.1. Role of spermidine in cell division ...... 9
1.2.2. Role of thermospermine in vascular development...... 10
1.3. References ...... 12
CHAPTER II. ALTERED EXPRESSION OF POLYAMINE TRANSPORTERS REVEALS A
ROLE FOR SPERMIDINE IN THE TIMIMG OF FLOWERING AND OTHER
DEVELOPMENTAL RESPONSE PATHWAYS ...... 21
2.1. Abstract…………………………………………...... 21
2.2. Introduction …………………………………………………………………… 22
2.3. Methods and Materials ...... 24
2.3.1. Plant material and growth conditions ...... 24
2.3.2. Phenotypic analysis ...... 25
2.3.3. Gene expression analysis ...... 26
2.3.4. GUS analysis ...... 26
2.3.5. Subcellular localization of PUTs ...... 26
2.3.6. Polyamine analysis...... 27
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2.3.7. Statistical analysis ...... 28
2.4. Results ...... 29
2.4.1. Promoter analysis of AtPUT5 ...... 29
2.4.2. Subcellular localization of PUTs by transient expression in N. benthamiana
...... 29
2.4.3. Phenotypic changes resulting from altered expression of PUTs ...... 29
2.4.4. Constitutive expression of PUTs delays senescence ...... 30
2.4.5. Effect of PUT expression on polyamine levels in rosette leaves ...... 31
2.4.6. Gibberellins suppress the delay of flowering in pPUT5:OsPUT1 plants 32
2.5. Discussion ...... 32
2.6. Acknowledgement ...... 37
2.7. Accepted Manuscript ...... 37
2.8. References ...... 39
CHAPTER III. THE EFFECT OF HIGH TEMPERATURE ON THE DELAY OF
FLOWERING IN PUT5 PLANTS… ...... ……………………. 54
3.1. Polyamines and Abiotic stresses ...... 54
3.1.1. High-Temperature Stress ...... 54
3.1.2. Low-Temperature Stress ...... 55
3.1.3. Drought Stress ...... 55
3.1.4. High Salinity Stress...... 56
3.2. High temperature suppresses the delay of flowering in PUT5:OsPUT1 plants . 57
3.3. References ...... 59
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CHAPTER IV. DUAL FUNCTIONING OF PLANT ARGINASES PROVIDES A THIRD
ROUTE FOR PUTRESCINE SYNTHESIS ...... ……………………………. 65
4.1. Abstract ...... 65
4.2. Introduction ...... 66
4.3. Materials and Methods ...... 68
4.3.1. Phyre2 analysis ...... 68
4.3.2. Phylogenetic analysis ...... 68
4.3.3. DNA sources and constructs ...... 68
4.3.4. Subcellular localization analysis ...... 69
4.3.5. Yeast complementation assays ...... 70
4.3.6. Growth rate of yeast strains without polyamine supplementation ...... 71
4.3.7. Agmatinase activity assays ...... 71
4.3.8. In vitro assay of ADC2 and ARGAH2 ...... 72
4.4. Results ...... 73
4.4.1. Plant arginases have agmatinase activity ...... 73
4.4.2. ADC2 and ARGAH2 together form a plastid putrescine pathway ...... 76
4.4.3. The plastid pathway is activated in stress responses ...... 76
4.4.4. Soybeans also have a complete plastid putrescine pathway, most plants do not ...... 77
4.5. Discussion ...... 78
4.6. Acknowledgements ...... 85
4.7. Accepted Manuscripts ...... 85
4.8. References ...... 87
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CHAPTER V. ATPDR11 IS A BROAD SUSBTRATE TRANSPORTER OF POLYAMINES
AND SELECTED AMINO ACIDS………………………………………………………… 112
5.1. Introduction ...... 112
5.2. Materials and Methods ...... 115
5.2.1. Phylogenetic analysis ...... 115
5.2.2. Plant material and growth conditions ...... 115
5.2.3. Phenotypic analysis ...... 115
5.2.4. Strains, media and reagents...... 115
5.2.5. DNA cloning ...... 116
5.2.6. Functional complementation assay ...... 116
5.2.7. Amino acid assay ...... 117
5.2.8. Time course dependent assay...... 117
5.3. Results ...... 118
5.3.1. Phylogenetic analysis of AtPDR11 ...... 118
5.3.2. The tissue/organ expression of AtPDR11 by microarray analysis ...... 119
5.3.3. Phenotypic characteristics of AtPDR11 mutant ...... 119
5.3.4. Involvement of AtPDR11 in abiotic stresses ...... 121
5.3.5. Functional characterization of AtPDR11 in yeast ...... 122
5.3.6. Heterologous expression of the AtPDR11 in yeast ...... 123
5.3.7. Transport of polyamines by AtPDR11 ...... 124
5.4. Discussion ...... 125
5.5. References ...... 127
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CHAPTER VI. FUNCTIONAL CHARACTERIZATION OF A VACUOLAR POLYAMINE
TRANSPORTER ...... ………………………………………………………… 134
6.1. Introduction ...... 134
6.2. Materials and Methods ...... 137
6.2.1. Plant materials and growth conditions ...... 137
6.2.2. Phenotypic analysis ...... 137
6.2.3. Polyamine toxicity assay...... 137
6.2.4. Subcellular localization of AtOCT5 ...... 138
6.3. Results ...... 138
6.3.1. Phenotypic characteristics of AtOCT5 mutant ...... 138
6.3.2. Involvement of AtOCT5 in abiotic stresses ...... 139
6.3.3. Polyamine toxicity assay in yeast ...... 140
6.3.4. Subcellular localization of ATOCT5 ...... 141
6.4. Discussion ...... 142
6.5. References ...... 145
CHAPTER VII. SUMMARY ...... ………………………………………………………… 149
7.1. Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways and the effect of high temperature on the delay of flowering in PUT5 plants ...... 149
7.2. Dual functioning of plant arginases provides a third route for putrescine synthesis
...... 150
7.3. PDR11 is a broad substrate transporter of polyamines and selected amino acids 151
7.4. Functional characterization of a vacuolar polyamine transporter ...... 152
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APPENDIX A. ARTICLE REPRINT PERMISSION ...... 153
xi
LIST OF FIGURES
Figure Page
1.1 Structure of major polyamines ...... 1
1.2 Factors affecting polyamine homeostasis ...... 2
1.3 Polyamine biosynthesis ...... 4
1.4 Polyamine catabolism ...... 6
1.5 Polyamine transporters in E. coli and S. cerevisiae ...... 7
1.6 Hypusination of eLF5A ...... 10
2.1 GUS-expression driven by the AtPUT5 promoter in A. thaliana ...... 46
2.2 Subcellular localization of AtPUT5 and OsPUT1 to the ER by transient expression in N.
benthamiana leaves ...... 47
2.3 Transient expression of AtPUT2, AtPUT3, and OsPUT3, tagged with GFP, in N.
benthamiana leaves ...... 48
2.4 Increased expression of PUTs results in larger leaves ...... 49
2.5 Increased expression of PUTs results in a delay of flowering ...... 49
2.6 A. thaliana plants with constitutive expression of OsPUT1 or OsPUT3 are delayed in
senescence ...... 50
2.7 Polyamine levels in rosette leaves of A. thaliana plants ...... 50
2.8 Gibberellins promote early flowering in pPUT5:OsPUT1 plants ...... 51
2.S1 Clustal alignment of OsPUT1 and PUT5...... 52
2.S2 Clustal alignment of chloroplast-localized PUTs ...... 52
2.S3 Expression analysis by RT-PCR ...... 53
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3.1 Increased expression of PUTs at 28 results in the disappearance of delay of flowering
phenotype ...... ℃ 58
4.1 Biosynthetic strategies for the synthesis of putrescine ...... 96
4.2 Plant agmatinases show conservation of key active site residues with agmatinase from
Deinococcus radiodurans ...... 96
4.3 Predicted 3D structure of AtARGAH2 is highly similar to the crystal structure of
Deinococcus radiodurans (DR) agmatinase ...... 97
4.4 Characterization of plant agmatinases by complementation in yeast ...... 98
4.5 Growth of yeast strains in the absence of putrescine ...... 99
4.6 HPLC analysis confirms agmatinase activity of AtARGAH2 ...... 99
4.7 HPLC analysis of arginine decarboxylase and arginase/agmatinase show accumulation
of agmatine and putrescine ...... 100
4.8 Confocal microscopy showing plastid localization of AtADC2 and AtARGAH2 in
transiently expressed N. benthamiana leaves ...... 101
4.9 In four stress responses of A. thaliana, ARGAH2 expression clusters with ADC2 ... 102
4.10 Confocal microscopy showing chloroplast localization of GmADC2 and GmARGAH in
transiently expressing N. benthamiana leaves ...... 103
4.S1 Maximum likelihood phylogeny of selected plant arginine ...... 104
4.S2 Maximum likelihood phylogeny of selected plant arginase/agmatinases ...... 105
4.S3 Fasta alignment file of plant species with Arginine decarboxylases ...... 106
4.S4 Fasta alignment file of plant arginase/agmatinases ...... 109
5.1 Topology of ABC transporters...... 113
5.2 Structure of paraquat ...... 114
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5.3 Phylogenetic analysis of AtPDR protein sequences...... 118
5.4 Predicted Tissue/Organ Expression of AtPDR11 ...... 119
5.5 Phenotypic Characteristics of wildtype and AtPDR11Δ plants ...... 120
5.6 Analysis of salt tolerance in wildtype and AtPDR11Δ plants ...... 121
5.7 Functional complementation of AtPDR11 in yeast mutant agp2Δ ...... 122
5.8 Amino acid transport assay of AtPDR11 in yeast mutant 22Δ8AA ...... 123
5.9 Time course uptake of [H3] spermidine ...... 124
6.1 Predicted membrane topology of OCT1 from rats and OCT5 from A.thaliana...... 136
6.2 Phenotypic Characteristics of wildtype and AtOCT5Δ plants...... 139
6.3 Analysis of salt tolerance in wild-type and AtOCT5Δ plants ...... 140
6.4 Polyamine toxicity assay in S. cerevisiae ...... 141
6.5 Sub-cellular localization of AtOCT5 ...... 142
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LIST OF TABLES
Table Page
2.1 Primer sequences used in this study...... 45
2.2 Phenotypic variation of A. thaliana genotypes ...... 45
3.1 Phenotypic variation of A. thaliana genotypes at 28 ...... 57
4.S1 Predicted plastid localization of arginine decarboxylase℃ genes from sequenced plant
genomes by PLPred, TargetP, Predotar and WolfPsort ...... 110
4.S2 Predicted plastid localization of arginase/agmatinase genes from sequenced plant
genomes by PLPred, TargetP, Predotar and WolfPsort ...... 111
4.S3 Primers used to amplify gene targets ...... 111
6.1 Structure comparison of putrescine, spermidine, spermine and metformin ...... 136
1
CHAPTER I.
INTRODUCTION
1.1. What Are Polyamines?
Polyamines are the oldest known substances in biochemistry. The history of polyamines dates back to about 300 years [1] and was first discovered by Antonie van Leeuwenhoek, who is also known as “the father microbiology”. Leeuwenhoek observed certain crystal formations in aging human sperm samples through his primitive microscope in 1670’s [2]. In 1920’s, Otto
Rosenheim determined the chemical composition and the structure of spermine [3]. Spermidine was also discovered and named at the same time. There are four major polyamines - the diamine putrescine (Put), triamine spermidine (Spd) and tetramine, spermine (Spm) and thermospermine
(tSpm)
Figure 1.1. Structure of major polyamines. 2
Polyamines (PAs) are low molecular weight small organic cations present in all living organisms [5]. They are known to be essential for the growth and development of prokaryotes and eukaryotes [6,7]. The major polyamines are found as free polyamines or as conjugated polyamines, forming conjugates with phenolic acids and other low molecular weight compounds resulting in soluble conjugates and with macromolecules such as proteins, phospholipids, nucleic acids forming insoluble conjugates. They affect the growth and development of a plant and play pivotal roles in cell division, fruit ripening, root formation, senescence, abiotic stress and infection by viruses and fungi [8, 9, 10]. Because of the various functions, the homeostasis of polyamines is vital and is assured by the regulation of biosynthesis, conjugation, catabolism, transport and excretion.
Figure 1.2. Factors affecting polyamine homeostasis.
1.1.1. Polyamine Biosynthesis
The polyamine biosynthetic pathways are conserved from bacteria to animals and plants
[6]. In plants, polyamines have been localized to the cytoplasm, vacuoles, mitochondria and chloroplasts [11]. The biosynthesis starts from two amino acid precursors – arginine and 3 methionine. There exist two biosynthetic pathways for polyamines. The first pathway involves the synthesis of putrescine from arginine via the conversion of arginine to ornithine by arginase and the conversion of ornithine to putrescine by ornithine decarboxylase (ODC). The second pathway involves the synthesis of putrescine from arginine by a three-step process catalyzed by arginine decarboxylase (ADC), agmatine iminohydrolase (AIH) and N-carbamoyl putrescine amidohydrolase (CPA).
Spermidine is made from putrescine by the action of spermidine synthase (SPDS). An aminopropyl group is transferred from the decarboxylated S-adenosylmethionine (dcSAM) for the synthesis of spermidine. Decarboxylated S-adenosylmethionine is synthesized from methionine by a two-step process catalyzed by methionine adenosyltransferase and S- adenosylmethionine decarboxylase (SAMDC). Spermine is made from spermidine via spermine synthase. Thermospermine is an isomer of spermine.
However, the model plant A. thaliana does not contain a gene coding for ornithine decarboxylase [12]. It has two genes coding for arginine decarboxylases (ADC1 and ADC2), [13,
14] and one each for agmatine iminohydrolase and N-carbamoyl putrescine amidohydrolase respective [15, 16]. For the synthesis of spermidine, it has two genes coding for spermidine synthase (SPDS1 and SPDS2) [17] and at least four genes coding for Decarboxylated S – adenosylmethionine (SAMDC1, SAMDC2, SAMDC3 and SAMDC4) [18, 19]. For the synthesis of spermine and thermospermine, it has two genes coding for spermine synthase and thermospermine synthase (SPMS and ACL5) [20, 21, 22].
. 4
Figure 1.3. Polyamine biosynthesis. Green arrows – plant pathways, blue arrows – bacterial pathways, red arrows – animal pathways. ADC – arginine decarboxylase, AIH – agmatine imunohydrolase, CPA – N-carbamoyl putrescine amidohydrolase, ODC – ornithine decarboxylase, SPDS – spermidine synthase, SPMS – spermine synthase, MAT - Methionine adenosyltransferase, AdoMetDC - S-adenosylmethionine decarboxylase.
1.1.2. Polyamine Catabolism
The enzyme polyamine oxidase (PAO) is a flavin adenine dinucleotide (FAD) containing an enzyme that uses N-acetyl derivatives as substrates [23]. In plants, Polyamine oxidases 5 catalyze the conversion of spermidine and spermine to 4-aminobutanal and N- (3-aminopropyl)-
4-aminobutanal respectively, with the production of 1, 3-diaminopropane and H2O2 [24, 25, 26].
The enzyme spermine oxidase (SMO) is an FAD-dependent amine oxidase which directs the back conversion of spermine to spermidine with the production of 3-aminopropanal and
H2O2; this was first identified in mammalian cells [27, 28, 29]. Fms1 was an enzyme with similar enzymatic activity found in yeast [30]. A. thaliana contains five polyamine oxidase-like genes [31]. Out of the five, AtPAO1 catalyzes the same reaction as spermine oxidase and FMS1
(fenpropimorph resistance multicopy suppressor 1), demonstrating the existence of back conversion of polyamine in plants [32]. Polyamine oxidase converting spermidine to putrescine needs further studies.
Diamine oxidase (DAO) is a copper-containing enzyme that catabolizes polyamines and prefers diamines as substrates [33, 34]. Diamine oxidase catalyzes the oxidation of putrescine to
4-aminobutanal with the production of NH3 and H2O2. The resulting 4-aminobutanal is further
aminobutyric acid via aldehyde dehydrogenase [35]. A. thaliana contains-ץ metabolized to twelve diamine oxidase-like genes [31]. Out of twelve, ATDAO1 is biochemically characterized
[36]. Functional analysis of the remaining 11 genes needs further studies. 6
Figure 1.4. Polyamine catabolism. a. Catabolism of putrescine. b. Catabolism of spermidine and spermine. Green arrows – plant pathways, blue arrows – bacterial pathways, red arrows – animal pathways. PAO – polyamine oxidase, DAO – diamine oxidase, SPMO – spermine oxidase.
1.1.3. Polyamine Transport
Polyamine transporters have been well characterized in microorganisms such as
Escherichia coli and Saccharomyces cerevisiae. There are four polyamine transport systems in 7
E.coli - spermidine-preferential uptake system (PotABCD), putrescine-specific uptake system
(PotFGHI), putrescine transport system (PotE), and cadaverine transport system (CadB) [37, 38].
Out of the four, PotABCD and PotFGHI are ATP-binding cassette transporters. There are nine polyamine transport proteins identified in S. cerevisiae - TPO1–5, UGA4, GAP1, DUR3, and
SAM3. TPO1–4, are efflux pumps for xenobiotics which also recognizes polyamines. TPO5 is localized to the post-Golgi complex and is an efflux pump for polyamines. UGA4 is localized to
aminobutyric acid. GAP, is localized to the-ץ the vacuoles and takes up putrescine along with cytoplasmic membrane and takes up polyamines into the cytoplasm along with amino acids.
DUR3 and SAM3, carry polyamines preferentially into the cytoplasm [38].
Figure 1.5. Polyamine transporters in E. coli and S. cerevisiae. a. Polyamine transporters in E. coli. b. Polyamine transporters in S. cerevisiae. Adopted from [39]. 8
In plants, the first published evidence of polyamine transporters revealed five polyamine uptake transporters (PUTs) from A. thaliana (AtPUT1-5) and three from O. sativa (OsPUT1-3)
[40]. Full lengths cDNAs were transformed into yeast agp2Δ mutant, which was deficient in high-affinity spermidine uptake [41]. The study identified OsPUT1, which complemented the agp2 mutant phenotype as spermidine specific transporter. Radiological uptake assays and competitive inhibition analyses in the yeast transformants revealed that OsPUT1 preferentially transported spermidine with K m = 15.2 μM [42]. The similar experimental approach was used to identify other polyamine transporters in rice and A. thaliana, including OsPUT2, OsPUT3,
AtPUT1, AtPUT2, and AtPUT3 [40].These polyamine transporters had affinities to both putrescine and spermidine, with Km values of 0.94–3.3 μM and 28.7–33.4 μM for spermidine and putrescine uptake, respectively.
In plants, it has been described that multidrug transporters are potential candidates for polyamine transport [43]. Paraquat being the structural analog of polyamines, share the same transport system and therefore paraquat transporters are suggested to transport polyamines as well [44, 45].
1.2. Polyamines and Development
The establishment of polyamine levels at different stages of plant development provides correlation of polyamines with cell division, flowering, and other developmental processes.
Studies have shown the occurrence of crosstalk between polyamines and hormones such as gibberellins, auxins, and ethylene. But the underlying mechanism for the interactions has not yet been established [46, 47, 4]. Genetic approaches have revealed the importance of polyamines in some developmental processes such as the involvement of spermidine in the posttranslational 9 modification of eIF5A, implications of HCAA conjugates in pollen development and the role of thermospermine in vascular tissue development.
1.2.1. Role of spermidine in cell division
Loss-of-function mutants of ODC (spe1), SAMDC (spe2) and SPDS (spe3) in S. cerevisiae were unable to grow in the absence of exogenous polyamines [48, 49, 50]. The growth of spe1, spe2, or spe3 yeast mutants could be restored by exogenous application of either
Spermidine or Spermine. Spermidine has been proved to be essential for cell division and spermine is known to be essential due to its oxidation to spermidine via FMS1 [51]. The importance of spermidine for cell survival has different possible explanations, related to the requirement of polyamines for essential cellular functions, such as the activation of the eukaryotic translation initiation factor eIF5A.
eIF5A is known to act on RNA and proteins, essential for the translation machinery or as initiation factor [52]. Activation of eIF5A takes place by hypusination of a specific lysine residue in its N-terminal domain. Hypusination of eIF5A is a post-translational modification that requires the activity of the deoxyhypusine synthase (DHS) enzyme. Deoxyhypusine synthase transfers the amino butyl moiety of spermidine to eIF5A to form the deoxyhypusine residue.
Hypusination is followed by the hydroxylation of deoxyhypusine to hypusine and is catalyzed by deoxyhypusine hydroxylase (DOHH) [53]. Hypusination of eIF5A is essential for cell division in eukaryotes and is dependent on the availability of free spermidine in the cytoplasm. In
A.thaliana, double loss of function mutants such as adc1 adc2, spds1 spds2, and samdc1 samdc4 are impaired in the biosynthesis of polyamines and are embryo lethal [54, 55, 19]. 10
Figure 1.6. Hypusination of eLF5A. DHS - deoxyhypusine synthase, DOHH - deoxyhypusine hydroxylase. Modified from klemkelab.ucsd.edu.
1.2.2. Role of thermospermine in vascular development
An ethyl methanesulfonate (EMS) mutagenesis screen revealed a mutant of acaulis5
(acl5), which affected the stem elongation in the Landsberg erecta (Ler) accession in A.thaliana
[56]. A similar phenotype was also seen in Columbia background suggesting that it is a background independent phenotype [57]. Allelic mutants of acaulis5 (acl5-1 and acl5-2), inhibited the elongation of the stem during the reproductive stage. acl5-1 mutants did not have any phenotypes before flowering and the phenotypes seen during reproductive stage could not be rescued by exogenous application of hormones. The vascular tissues in the stem of acl5-1 11 mutants have over the proliferation of the xylem elements and do not develop xylem fibers resulting in cell death in developing xylem vessel elements [56, 58]. Phenotypes seen in the mutants of acl5 have been used to identify pathways involved in thermospermine signaling.
In bacterial cultures, recombinant ACL5 could convert spermidine to spermine and therefore ACL5 was originally known as spermine synthase (SPMS) [20]. It was later known that ACL5 might have other enzymatic activity than spermine biosynthesis. The mutants of spms-1 were almost depleted in spermine levels but had no phenotypic differences from the wild type. The mutants of acl5 did not have any differences in spermine levels. The spms-1 acl5 double mutants had acl5 phenotypes. Characterization of a thermospermine synthase from the diatom Thalassiosira pseudonana that is a homolog of A. thaliana ACL5 revealed thermospermine as the product of ACL5 reaction [59].The identity of ACL5 as a thermospermine synthase was confirmed in the absence of thermospermine in acl5 mutant by exogenous application of thermospermine [60].
12
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20
CHAPTER II.
ALTERED EXPRESSION OF POLYAMINE TRANSPORTERS REVEALS A ROLE
FOR SPERMIDINE IN THE TIMING OF FLOWERING AND OTHER
DEVELOPMENTAL RESPONSE PATHWAYS
2.1. Abstract 21
Changes in the levels of polyamines are correlated with the activation or repression of developmental response pathways, but the role of polyamine transporters in the regulation of polyamine homeostasis and thus indirectly gene expression, has not been previously addressed.
Here we show that the A. thaliana and rice transporters AtPUT5 and OsPUT1 were localized to the ER, while the AtPUT2, AtPUT3, and OsPUT3 were localized to the chloroplast by transient expression in N. benthamiana. A. thaliana plants that were transformed with OsPUT1 under the control the PUT5 promoter were delayed in flowering by 16 days. In contrast, put5 mutants flowered four days earlier than WT plants. The delay of flowering was associated with significantly higher levels of spermidine and spermidine conjugates in the leaves prior to flowering. A similar delay in flowering was also noted in transgenic lines with constitutive expression of either OsPUT1 or OsPUT3. All three transgenic lines had larger rosette leaves, thicker flowering stems, and produced more siliques than wild type plants. In contrast, put5 plants had smaller leaves, thinner flowering stems, and produced fewer siliques. Constitutive expression of PUTs was also associated with an extreme delay in both plant senescence and maturation rate of siliques. These experiments provide the first genetic evidence of polyamine transport in the timing of flowering, and indicate the importance of polyamine transporters in the regulation of flowering and senescence pathways.
2.2. Introduction
The polyamines putrescine, spermidine and spermine were amongst the first metabolites to be isolated from living organisms [1] but how these compounds function at a molecular level is largely uncharted territory. Polyamines have hormone-like properties in that changes in polyamine levels are associated with fundamental physiological processes such as embryogenesis, flowering, fruit development and ripening, senescence, and tissue responses to 22 biotic and abiotic stresses [2]; but the levels of polyamines are much higher than true hormones such as auxins, gibberellins, or brassinosteroids. In A. thaliana, putrescine levels range from 10-
50 nM/g FW, while spermidine levels are typically higher (50-80 n /g FW) and spermine levels fall in the range of 10-50 nmol /gFW [3, 4]. The identification of thermospermine in plant tissues is a more recent development, largely because this compound is not easily resolved from spermine [4]. Levels of thermospermine in different tissues are estimated to be in the range of
0.5 to 2 nM/g FW. Conjugated forms of polyamines that include alkaloids, and caffeic, coumaric, and ferulic acids form the largest pools of polyamines in plants [5-7]. More recent evidence suggests that some phenolamides have specific roles such as pollen development [2] and defense responses to pathogens and insects [6]. The extent of back conversion of such compounds to the polyamines is also unknown [8].
These estimates of pool sizes may be misleading, as polyamines could be concentrated in cellular compartments or bound to macromolecules. The association of polyamines with other macromolecules has been estimated for spermidine and spermine for mammalian cells, and putrescine and spermidine in E. coli [9]. While polyamines were found not to have a strong affinity for cytosolic proteins, a significant fraction of polyamines were bound to RNA. In bovine lymphocytes, 57% of the spermidine pool, and 65% of spermine was bound to RNA, while in E. coli, 48% of the putrescine pool, and 90% of the spermidine was bound to the RNA.
In mammalian cells, the percentage of spermidine bound to DNA, ATP and phospholipids was
13%, 12%, and 3%, and that of spermine was 18%, 9% and 2.5%, respectively. Importantly, the levels of “free” spermidine and spermine in mammalian cells was estimated to be 15% and 5% respectively of the total pools of these polyamines [9]. Putrescine levels were not estimated in mammalian cells. However, in E. coli, putrescine was found to have a lower affinity for other 23 macromolecules than the larger polyamines. In those cells, the amount of free putrescine was estimated to be 39% of the total pool, while only 4% of the total amount of spermidine in the cell was estimated to be free. In plants, non-aqueous fractionation has been used to fractionate metabolites into a cytosolic compartment, vacuolar, and chloroplast fractions [10]. In this study
61% of the spermidine and 45% of soluble putrescine were estimated to be localized with the chloroplast fraction, while the cytosolic fractions contained 33% and 16% respectively of the putrescine and spermidine pools.
A role of polyamines as an additional regulator of flowering has been suggested from both feeding experiments and transgenic manipulation of polyamine biosynthesis. In Sinapsis alba, the transition to flowering was marked by changes in the levels of polyamines and polyamine conjugates in xylem and phloem exudates [11]. Furthermore, the inhibition of flowering by the application of the ornithine decarboxylase inhibitor DFMO to the leaves and its reversal by application of putrescine to the roots, suggested that putrescine regulated the timing of flowering. In A. thaliana, feeding spermidine via the roots under permissive flowering conditions resulted in a delay of flowering [12]. Overexpression of arginine decarboxylase resulted in the accumulation of putrescine in the leaves, and plants that were both dwarfed and delayed in flowering [13]. The dwarfed phenotype suggested that high putrescine levels might be inhibiting either gibberellic acid (GA) synthesis, or the response of leaf tissues to GA. Both the dwarfing phenotype, and the delay in flowering were alleviated by spraying leaves with GA.
In prior work, we identified a clade of Polyamine Uptake Transporters (PUTs) that act as spermidine-preferential transporters [14, 15]. To demonstrate that transport is an essential component of polyamine homeostasis, we first identified PUTs that were altered in their subcellular location in plant leaves. Then, we tested the hypothesis that alterations in the 24 expression of transporters localized to those membranes would result in changes of plant phenotypes. Here, we show that the A. thaliana transporter PUT5 (At3G19553) is localized to the ER, while PUT2 (At1G31830) and PUT3 (At5G05630) are localized to the chloroplast. In plants with heterologous expression of OsPUT1 (localized to the ER) or OsPUT3 (localized to the chloroplast) we noted an accumulation of spermidine and spermidine conjugates in the leaves prior to the onset of flowering, a delay in the timing of flowering, larger leaves, thicker stems, and more flowers. In contrast, put5 plants had smaller leaves, flowered earlier than WT plants, had thinner stems, and produced fewer flowers.
2.3. Materials and Methods
2.3.1. Plant material and growth conditions
Homozygous mutants of put5 (Salk_007135) were obtained from the Arabidopsis
Biological Resource Center, Columbus, OH. Plants were grown at 22°C under LD conditions
(16 h light/8 h dark). Rice clones OsPUT1 (AK068055) and OsPUT3 (AK070314) were obtained from the Rice Genome Resource Center (http://www.rgrc.dna.affrc.go.jp/index.html). 25
The full-length coding sequences of OsPUT1 and OsPUT3 were amplified by PCR using the primer sequences listed in Table 1 and cloned into the pENTR/D-TOPO cloning vector
(Invitrogen, Carlsbad, USA) according to the supplier’s protocol. The resulting plasmids were used to mobilize target genes into the plant expression vector, pGWB2 [16] to generate pGWB2-
OsPUT1 and pGWB2-OsPUT3 by LR recombination reaction (Invitrogen, Carlsbad, USA). The plant expression vectors were then transformed into Agrobacterium tumefaciens strain GV3101 followed by a transformation into Arabidopsis Col-0 plants using the floral dip method [17].
Transgenic plants were screened by germination of seeds on ½ MS plates containing 100 µg/ml of kanamycin. Seeds from 10 independent T3 lines for each of OsPUT1 and OsPUT3 were pooled and used for further analysis. Transgenic plants expressing pPUT5:OsPUT1 were generated by cloning and transforming, the 5’ upstream region (1.3 kb) of At3g19553 (see Table
1) followed by the full-length coding sequence of OsPUT1 in pEG301 [18] using the same steps as described above. These plants were selected by spraying with BASTA. Seeds from 12 plants after T3 were pooled as all plants were phenotypically similar, and used in subsequent experiments.
2.3.2. Phenotypic analysis
Seeds were surface sterilized, vernalized at 4°C for 3 days and planted in soil. Plants were grown in a growth chamber at 22°C and a relative humidity of 55%. Stem thickness was measured by using a vernier caliper (Fowler tools and instruments, Boston, USA). Fluorescence measurements were made using a FluorPen FP 100 (Photon System Instruments) [19]. Flowering time was measured as the day when the floral stem was one cm above the rosette. To test the effect of GA on flowering time, the plants were sprayed four times with 100 µM GA, at four day intervals starting at eight days after planting. 26
2.3.3. Gene expression analysis
Total RNAs were extracted from leaf tissues using RNase easy plant mini kit (Qiagen,
CA). The purity and concentration was measured using the Nanodrop 2000 (Thermo Fisher, DE).
Gel electrophoresis was used to ascertain the integrity of the RNA. cDNA synthesis was performed using SuperScript® III First-Strand Synthesis SuperMix and oligo(dT) primers
(Invitrogen, CA) following the manufacturer’s protocol. Gene expression analyses were performed via RT-PCR using Eppendorf™ Mastercycler™ pro PCR System. All primers used are listed in Table 1. PCR was performed using Taq 2X Master Mix (NEB, MA). PCR conditions were 20 cycles of 95°C for 20 s, 55°C for 30 s, 72°C for 1 min, with final extension of
72°C for 5 min. The PCR amplicons were assessed on 1% agarose gel electrophoresis.
2.3.4. GUS analysis
The 5’ upstream region of AtPUT5, 1320 bp in length, was amplified by PCR. The amplified product was cloned into the Gateway promoter analysis vector pBGWFS7 by Gateway technology [20]. The resulting construct was then transformed in A. thaliana (Col-0) by floral dip method [17]. BASTA resistant seedlings were selected, transferred to soil and grown at 22
C in a growth chamber. Histochemical staining of GUS expression from eight resistant lines was⁰ performed by infiltration with X-Gluc buffer (2 mMX-Gluc, 50 mM NaPO4 pH 7.0, Triton-X
(0.5%), 0.5 mM K-ferricyanide, 0.5 mM K-ferrocyanide) [21].
2.3.5. Subcellular Localization of PUTs
Full length sequences of AtPUT5, AtPUT2 (At1G31820); AtPUT3 (At5G05630);
OsPUT1 and OsPUT3.1 were cloned into the GFP expression vectors pGWB6 or pGWB5 using the GATEWAY® recombination system [16]. Inserts were verified by sequencing, and vectors 27 were transformed into Agrobacterium tumefaciens strain GV3101. [22]. The mCherry-ER
Marker (ER-rbCD3960) [23] was obtained from ABRC and used as an organellar marker.
Images of one micron sections through the leaf were acquired using a Leica TCS SP5 multi- photon laser scanning confocal microscope at 12-72 hours after infiltration. Samples were imaged using a Leica TCS SP5 laser scanning confocal microscope (Leica Microsystems,
Bannockburn, IL) and the Leica Application Suite Advanced Fluorescence (LASAF) program.
Images were acquired in the XYZ plane in 1 µm steps with a 63X oil objective (NA 1.40) using the sequential scan mode to eliminate any spectral overlap in the individual fluorophores.
Specifically, GFP was excited at 488 nm and detected at 510 nm. The mCherry was excited at
561nm and detected at 610 nm. Plastids were excited at 633nm and detected at 670 nm. GFP signals were false-colored green and mCherry signals were false-colored red. Background fluorescence from untransformed leaves of plants at similar laser excitation settings were acquired and subtracted from images to identify fluorescence generated by tagged proteins.
Images were merged using ImageJ [24]. Images were acquired from multiple leaf sections at different time points following infiltration on at least two separate occasions for each construct.
2.3.6. Polyamine analysis
Leaves of Arabidopsis plants were extracted in 5% cold perchloric acid (300 mg fresh weight/ml) in an ice bath. Plant extracts were centrifuged at 10,000 g for 10 minutes. The supernatant, containing the soluble polyamines and hydrolysable conjugates, was stored at -
20°C. The pellet containing insoluble conjugates was re-suspended in 1 M NaOH (300 mg fresh weight/ml) and stored at -20°C until dansylation. Soluble conjugated polyamines and insoluble conjugated polyamines were released from the supernatant and pellet fractions respectively, by hydrolysis with equal volumes of 12 M HCl at 100 C for 20 h. Dansyl derivatization was 28 performed as described (Marce et al., 1995). 200 µl of supernatant was mixed with an internal standard (40 µl of 0.05 mM 1,7-heptanediamine, Sigma-Aldrich), 200 l of saturated sodium carbonate (13%, w/v) and 400 µl of dansyl chloride in acetone (10 mg/ml). After vortexing, the mixture was incubated overnight at room temperature in the dark. Excess dansyl chloride was removed by adding 100 µl of L-proline (100 mg/ml) followed by an incubation for 30 min at room temperature in the dark. The dansylated polyamines were further extracted with 500 µl of toluene. The extraction was repeated two times. The organic phase containing polyamines was vacuum evaporated, and the residue was dissolved in 500 µl of methanol. The dansylated polyamines were separated and quantified by HPLC with an Agilent Technologies 1120 series
HPLC and Applied Biosciences 980 programmable fluorescence detector (excitation at 340 nm, emission at 500 nm). A Gemini® 5 μm C18 110 Å, 250 × 4.6 mm LC column (Phenomenex,
Torrance, CA) was used for the separation of polyamines. Dansylated polyamines were separated with the HPLC method of Smith et al., (1985) with modifications. Samples were eluted from the column with a programmed water: methanol solvent gradient over 40 minutes. The initial conditions were 10% methanol and 90% water pumping at a flow rate of 0.75 ml/min. The methanol concentration was increased to 60% over 4 minutes and then up to 80% over 11 minutes. Then the concentration of methanol was increased to 95% over 17 minutes. A standard curve was generated by measuring known amounts of polyamines and used for the estimation of polyamine content.
2.3.7. Statistical analysis
All data analyses were performed using the R Studio software program, V. 0.98.1062. A one-way T test was applied, and the difference between means were considered to be significant for P< 0.05. 29
2.4. Results
2.4.1. Promoter analysis of AtPUT5
Microarray analysis of PUT5 indicates that this gene shows highest expression in mature leaves, the cauline leaf, and the stem [25]. To determine the cellular distribution of this transporter in plant tissues, we fused the upstream sequence of PUT5 to GUS, and transformed these constructs into A. thaliana. At two and four weeks, the highest level of staining was observed in the major veins of the leaves (Fig. 1). At six weeks, staining was observed throughout the leaf, at the tops of flower petals, and the root shoot junction. However, in our constructs we did not note any expression in the stems.
2.4.2. Subcellular localization of PUTs by transient expression in N. benthamiana
Confocal analysis of GFP-AtPUT5 and GFP-OsPUT1 fusion constructs revealed a compartmental localization within the cytoplasm. Co-expression with an mCherry ER-marker indicated that both AtPUT5 and OsPUT1 were localized to the ER (Fig. 2). In contrast, confocal analysis of GFP fusions of AtPUT2, AtPUT3, and OsPUT3 showed that these PUTs were localized to the chloroplast in the leaves of N. benthamiana (Fig. 3).
2.4.3. Phenotypic changes resulting from altered expression of PUTs
The rice gene OsPUT1 was the first characterized member of the PUT family [14] and shares 56% identity and 71% homology with PUT5 (Fig.S1). Since both AtPUT5 and OsPUT1 are localized to the ER, put5 plants and A. thaliana plants transformed with pPUT5:OsPUT1 and p35S:OsPUT1 were used to assess how variation in the expression of an ER-localized polyamine transporter might affect plant phenotype. Plants with PUT5:OsPUT1 or p35S:OsPUT1 constructs enabled us to examine the consequences of tissue-specific vs constitutive changes in 30 gene expression on plant phenotypes. The top BLAST hits for the rice gene OsPUT3 rice gene in the A. thaliana genome are to PUT2 and PUT3. OsPUT3 shares 64% identity and 76% similarity to PUT2 and 62% identity and 77% similarity to PUT3 (Fig. S2). Thus, transformation of plants with the construct p35S:OsPUT3 was considered the best choice to evaluate the effect of overexpression of a plastid-localized polyamine transporter on plant development. To confirm expression of transgenes in transformant lines, and the absence of expression of PUT5 in put5 plants, we did RT-PCR of mRNA from rosette leaves (Fig. S3).
The phenotypes of the three overexpression lines were compared with wild type (WT) and put5 plants. Under LDs at 22oC, put5 flowered 4 days earlier than WT, which flowered at 27
± 3 days (Table 2). All of the overexpression lines were significantly delayed in flowering.
Plant expressing pPUT5:OsPUT1 flowered at 43 ±3 days, while plants expressing p35S:OsPUT1 and p35S:OsPUT3 flowered at 38 ±3 and 44 ±4 days respectively. The number of rosette leaves was positively correlated with time of flowering. At five weeks, the rosette leaves of the put5 were smaller than WT, while the leaves of three overexpression lines were two-fold larger than
WT leaves (Fig. 4 and Fig. 5). We also noted that overexpression lines had thicker and more rigid floral stems (Table 2, Fig. 4). The stems of the put5 plants near the base of the rosette were slightly thinner (0.74 mm) than WT plants (1.16 mm) but the floral stems in the overexpression lines ranged from 1.7 to 1.9 mm.
2.4.4. Constitutive expression of PUTs delays senescence
Constitutive expression of PUTs was also associated with an extreme delay in senescence. At 97 days, rosette leaves of p35S:OsPUT1 and p35S:OsPUT3 were still green, and had some photosynthetic electron transport capacity as assessed by Fv/Fm (Fig 6) compared to 31
WT plants. The siliques of these transformants also remained green and dessicated much more slowly. However, no delay of senescence relative to WT plants was noted for pPUT5:OsPUT1 plants.
2.4.5. Effect of PUT expression on polyamine levels in rosette leaves
Polyamine levels in leaves fluctuated during different developmental stages of the plant.
Polyamine levels were assessed at two weeks, four weeks, and six weeks to correspond with three periods in the life cycle of WT plants. At two weeks, the WT plants were at the early stages of development with no floral stems. At four weeks, WT plants were at the flowering stage, and at six weeks, WT plants had a large number of dried siliques. At week two in WT plants, pools of putrescine and spermine conjugates were less than 10 nmoles/g FW and did not change significantly over the three time points. Levels of spermidine conjugates were 34 ±2 nmoles/g
FW at two weeks and then declined to 15 ±3 and 9 ±2 nmoles/g FW at four and six weeks respectively (Fig. 7). While the phenotypes of put5 plants were distinguishable from WT plants, there were no significant differences from WT plants in any of the polyamine pools (Fig 7). In contrast, at two weeks, each of the overexpression lines had significantly higher levels of spermine, spermidine and spermidine conjugates than WT plants. pPUT5:OsPUT1 plants had significantly higher putrescine levels than WT plants. At four weeks, when WT were already flowering (Fig. 7) spermidine and spermidine conjugate levels, were not significantly different in pPUT5:OsPUT1 plants from WT plants, but they remained significantly higher in both of the
CaMv overexpression lines. At six weeks, the two lines with constitutively expressed PUTs had significantly higher levels of putrescine, spermidine and spermidine conjugates. The elevated levels of substrates in these lines were correlated with a delay in senescence that was not seen in pPUT5:OsPUT1 plants. 32
2.4.6. Gibberellins suppress the delay of flowering in pPUT5:OsPUT1 plants
Previous studies have shown that overexpression of arginine decarboxylase (the first step in the putrescine synthesis pathway) resulted in plants that accumulated putrescine in the leaves, and exhibited a delay of flowering, and a dwarf phenotype [13]. In contrast, plants with the pPUT5:OsPUT1 construct had significantly higher levels of putrescine than the WT at two weeks. To test whether the delay of flowering in pPUT5:OsPUT1 constructs could be alleviated by GA, leaves of these plants were sprayed four times with GA. At the time, when most of the put5 plants were starting to flower, all of the floral stems of GA-treated plants were more than 10 cm high. Thus, spraying the leaves of these plants enabled them to flower earlier than both put5 and WT plants, and more than two weeks earlier than expected (Fig. 8 and Table 1).
2.5. Discussion
This study provides the first experimental evidence that the regulation of polyamine transport plays a key role in both polyamine homeostasis and the downstream consequences of changes in cellular levels of polyamines and polyamine conjugates. The polyamine transporter mutant put5 flowered slightly earlier than WT plants, while flowering was delayed by 16 days in pPUT5:OsPUT1 plants. The delay of flowering was also noted in the constitutively expressed transgenic lines. The profiles of polyamine levels were unique for each of the three transgenic lines. However, a common feature that correlated with the phenotype of delayed flowering, was significantly higher leaf levels of both spermidine and spermidine conjugates at two weeks, and higher levels of spermidine conjugates in the rosette leaves at four weeks, in the two constitutively overexpressed lines.
PUT transporters have been characterized as unidirectional, preferential transporters of 33 spermidine [14, 15]. Here, we noted that OsPUT1 and AtPUT5 were localized to the ER, when transiently expressed in the leaves of N. benthamiana. Localization of AtPUT5/LAT3 to the ER by transfection of A. thaliana protoplasts was also reported [26]. While we used transient expression analysis in N. benthamiana of full length cDNAs of AtPUT2, AtPUT3, and OsPUT3, to localize these genes to the chloroplast, other groups have examined the localization of these genes in other plant tissues [26-29]. Confocal microscopy of AtPUT3/RMV1-GFP fusions in transgenic A. thaliana showed this protein to be localized to the plasma membrane in root cells
[27, 29]. Callus cultures of lhr1/put3 plants were also reduced in their ability to take up 2 µM spermidine relative to WT cells, consistent with the localization of this transporter to the plasma membrane in those tissues [29]. In stably transformed A. thaliana, GFP ATPUT2/PAR1-GFP fusions were localized to a subcellular compartment of roots cells, that in protoplasts prepared from leaf tissues was found to be co-localized with an ER marker [26]. The same research group reported that OsPAR1-GFP fusions were localized to the Golgi in both rice and A. thaliana protoplasts. There are many potential explanations that may account for the dual localization of proteins. For example, there are many proteins that are targeted to the chloroplasts or mitochondria in different tissues despite the striking differences in the import apparatus of these two organelles [30]. The plant glutamate receptor AtGRL3.4 was localized to both the plasma membrane and chloroplast using both a specific antibody and YFP-tagged fusion constructs [31].
The dual targeting of this protein was observed in WT plants, transiently expressed protoplasts of
A. thaliana and agroinfiltrated tobacco leaves. Some proteins such as the rice alpha amylase protein, and nucleotide pyrophosphatase/phosphodiesterases [32] are targeted to the plastid after first passing through the Golgi [33]. Protoplasts have been a valuable resource for localization studies, but protoplasts prepared from leaf tissues are not identical cell types, and the localization 34 of some proteins is both tissue and cell-type dependent [34]. Taken together, the localization experiments suggest that these transporters may, like other proteins be localized to more than one membrane under different conditions. This conclusion needs to be verified independently by immuno-localization experiments.
However, the in vivo experiments using detached leaves or isolated chloroplasts, directly supports the chloroplast localization of PUT2 and PUT3 in these tissues. The leaves of overexpression constructs of PUT3/RMV1 were found to be hypersensitive to paraquat ([27], Fig
3) a phenotype that would also be expected for a chloroplast-localized paraquat transporter.
Similarly, Li et al (2013) reported that paraquat accumulation in the chloroplast was markedly increased in overexpressing PAR1/PUT2 plants ([26]Fig. 6B). Both groups also noted that null mutations did not prevent the paraquat sensitivity of chloroplasts, a finding that is also consistent with our observations that both AtPUT2 and AtPUT3 are localized to the chloroplast.
It is interesting to note that the overexpression lines all had delayed flowering times, larger leaves and thicker stems, although the subcellular distribution of AtPUT5 and OsPUT1 is different from that of OsPUT2, AtPUT3, and OsPUT3. The direction of substrate transport for these proteins has not been clearly established, but the increased sensitivity of detached leaves to paraquat in transgenic lines overexpressing PUT2/RMV1 [27] or PUT3/PAR1 [26] is consistent with their localization to the chloroplast, and that they function as importers. We also have not yet demonstrated that proteins localized to the ER are functionally active, but the shared phenotypes of the overexpression lines of PUT5 and OsPUT1 with those of plants overexpressing OsPUT3, could mean that all transporters act to sequester spermidine from the cytoplasm. 35
Polyamines are not often referenced as significant regulators of flowering in plants [35,
36]. However a premature flowering phenotype which was also correlated with a significant increase in putrescine levels was recently reported [37]. The increase in putrescine levels was observed in plants that were deficient or silenced in N2-acetylornithine deacetylase which converts acetylornithine to ornithine, but the mutant plants also had lower seed production. In contrast, we observed increased levels of putrescine in pPUT5:OsPUT1 plants at two weeks, but these plants exhibited a delay in flowering (Fig. 6). A delay of flowering has also been reported in A. thaliana that were supplemented with spermidine via the roots [12]. Flowering was also delayed in transgenic lines overexpressing AtADC2 [13]. The dwarf phenotype of those plants suggested that the phenotype might be caused by an inhibition of gibberellin biosynthesis. Down regulation of genes in the gibberellin biosynthetic pathway was noted in the dwarf plants, and gibberellin treatment of the leaves reversed the inhibition of flowering [13]. By four weeks, the rosette leaves of all three transgenic lines were larger than WT plants, which would not be expected if the accumulation of spermidine or spermidine conjugates resulted in an inhibition of gibberellin synthesis. However, the application of gibberellins clearly promoted the onset of flowering, and these plants flowered even earlier than put5 plants. We therefore hypothesize that the inhibition of flowering due to the higher levels of spermidine and spermidine conjugates in young rosette leaves, is upstream of the activation of flowering due to the gibberellin response pathway. Sustained levels of spermidine or spermidine conjugates in the rosette leaves at four weeks appear not to be needed to cause the delay in flowering, as polyamine levels in pPUT5:OsPUT1 plants were not significantly different from WT plants. Yet these plants flowered later than p35S:OsPUT1 plants and at the same time as p35S:OsPUT3 plants. 36
The three overexpressor lines described here displayed a common set of other phenotypes, notably, larger leaves, and thicker stems. Taken as a whole, these developmental changes suggest the involvement of several regulatory pathways. One possibility is that spermidine and or spermidine conjugates are acting as signaling molecules. Changes in the levels of these signaling molecules are sensed by the cell and serve as a signal to activate hormone responsive pathways [38]. Plants overexpressing ADC2 were found to have increased tolerance to both freezing and drought stress [3, 39]. In these plants, microarray analysis pointed to altered expression of genes involved in auxin synthesis, auxin transport, auxin-related genes, along with genes associated with abiotic stress in plants [38]. Gene expression in hormone-responsive pathways were also affected. These included ABA, jasmonic acid, and ethylene pathways.
The thinner stems that we noted in put5 plants, and the thicker, more robust stems that are a feature of all of the overexpression lines of PUTs, may be related to alterations in thermospermine levels. Thermospermine is synthesized by ACL5 in vascular tissues [40] and vascular development occurs as a consequence of a feedback loop between auxin levels, HD-
ZIPIII genes and thermospermine [41]. Auxin promotes the synthesis of both ACL5 and BUD2
[42], genes that are needed for thermospermine synthesis. Thermospermine in turn, promotes the expression of SAC51 and other bHLH transcription factors, that together act as negative regulators of auxin signaling and perivascular divisions [43]. Increased expression of a cotton
ACL5 in A. thaliana resulted in taller plants [44] while silencing of the gene resulted in a severe dwarf phenotype [40]. Spermidine is a precursor to thermospermine, and increased levels of spermidine in stem tissues may result in increased synthesis of thermospermine, and thus taller plants. Changes in the expression of PUT transporters may also affect the redistribution of thermospermine. The phenotypes produced by enhanced PUT transport are consistent with an 37 overall increase in the expression of genes in the auxin, HD-ZIPIII, and thermospermine regulatory loop described above. The role of spermidine and spermidine transport in promoting vascular development is presently under investigation.
One of the great challenges for plant scientists is to be able to explain the phenotypic diversity within a single plant species. Here, we have shown that mutations that change the expression of a single spermidine transporter have pleiotropic effects. Thus, transporters that regulate the movement of hormone and other signaling compounds could provide a mechanism for generating phenotypic variation across species.
2.6. Acknowledgments
We thank Gerald Jones for the loan of a HPLC fluorometer and integrator used in polyamine analysis, Rachel Wilson for assistance with RNA extractions, and the Arabidopsis Stock Center for providing seeds of T-DNA insertion mutants and cDNA clones. This work was partially financed with support from BGSU Graduate College, a BGSU “Building Strength” grant in aid of research, and USDA-NIFA 2011-68004-30104.
2.7. Accepted Manuscript
Title: Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways.
Author: Sheaza Ahmed Menaka Ariyaratne Jigar Patel
Alexander E Howard Andrea Kalinoski Vipaporn Phuntumart
Paul F. Morris 38
PII: S0168-9452(16)30374-0
DOI: http://dx.doi.org/doi:10.1016/j.plantsci.2016.12.002
Reference: PSL 9530
To appear in: Plant Science
Received date: 15-9-2016
Revised date: 1-12-2016
Accepted date: 8-12-2016
Please cite this article as: Sheaza Ahmed, Menaka Ariyaratne, Jigar Patel, Alexander E Howard,
Andrea Kalinoski, Vipaporn Phuntumart, Paul F.Morris, Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways., Plant Science http://dx.doi.org/10.1016/j.plantsci.2016.12.002
39
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45
Table 2.1. Primer sequences used in this study.
Table 2.2. Phenotypic variation of A. thaliana genotypes.
46
Figure 2.1. GUS-expression driven by the AtPUT5 promoter in A. thaliana. (A) In two-week rosette leaves, staining is observed in the veins; (B) At four weeks, staining restricted to the major vein of rosette leaves; (C) At six weeks, staining is visible throughout the leaf blade; (D)
Staining is observed at the tip of the flower petals. (E) Staining is visible at the base of leaf petiole.
47
Figure 2.2. Subcellular localization of AtPUT5 and OsPUT1 to the ER by transient expression in N. benthamiana leaves. (A) Expression of GFP-AtPUT5 (B) Localization of mCherryER marker (C) Merged images showing co-localization of GFP and mCherry markers
(yellow). (D) Expression of GFP-OsPUT1 (E) Localization of mCherry-ER marker (F) Merged images showing co-localization of GFP and mCherry markers. Z-stack images were generated with Image J [24].
48
Figure 2.3. Transient expression of AtPUT2, AtPUT3, and OsPUT3, tagged with GFP, in N. benthamiana leaves indicate that these transporters are localized to the chloroplast. (A)
Expression of GFP-AtPUT2 (B) Chlorophyll autofluorescence (excitation 633 nm, emission 670 nm) (C) Merged images showing GFP localization and chlorophyll autofluorescence (yellow).
(D). Expression of GFP-AtPUT3; (E) chlorophyll autofluorescence; (F) Merged images showing
GFP localization and chlorophyll autofluorescence (yellow); (G) Expression of OsPUT3 (H) chlorophyll autofluorescence; (I) Merged images showing GFP localization and chlorophyll autofluorescence (yellow). Z-stack images were generated with Image J [24].
49
Figure 2.4. Increased expression of PUTs results in larger leaves. Rosette leaves of plants at
27 days. Note the reduction in petiole length in the overexpression lines.
Figure 2.5. Increased expression of PUTs results in a delay of flowering. Photograph of A. thaliana plants at six weeks. Note the thicker stems in the overexpressed lines. 50
Figure 2.6. A. thaliana plants with constitutive expression of OsPUT1 or OsPUT3 are delayed in senescence. Fluorescent measurements of the rosette leaves of p35S:OsPUT1 and p35S:OsPUT3 indicated that they were still photosynthetically active at 97 days.
51
Figure 2.7. Polyamine levels in rosette leaves of A. thaliana plants. Soluble (S), soluble conjugated (SC) and insoluble conjugated (IC) polyamines at two weeks (A) four weeks (B) and
6 weeks after planting(C). * indicates a significant difference from the WT pool at that time period.
Figure 2.8. Gibberellins promote early flowering in pPUT5:OsPUT1 plants. Plants expressing pPUT5:OsPUT1 that were sprayed with gibberellins flowered slightly ahead of put5 plants. 52
Figure 2.S1. Clustal alignment of OsPUT1 and PUT5 (At3g19553).
Figure 2.S2. Clustal alignment of chloroplast-localized PUTs. Full length sequences of
OsPUT3 is aligned with ATPUT2 (AT3G31830) and PUT3 (At5g05630). 53
Figure 2.S3. Expression analysis by RT-PCR. A. Lane 1: 100 bp DNA ladder; Lane 2:
Negative control; Lane 3: expression of OsPUT1 in p35S:OsPUT1; Lane 4: expression of
OsPUT1 in pPUT5:OsPUT1; Lane 5: WT, no expression of OsPUT1 in A. thaliana. Lane 6:
Negative control; Lane 7: expression of OsPUT3 in p35S:OsPUT3 plant, Lane 8: WT: no expression of OsPUT3 in WT A. thaliana; Lane 9: negative control: Lane 10: expression of
PUT5 in WT plants; Lane 11: No expression of PUT5 in put5 mutants. B. Agarose gel showing expression of the actin gene (At3G18780) as internal control.
54
CHAPTER III.
THE EFFECT OF HIGH TEMPERATURE ON THE DELAY OF FLOWERING IN
PUT5 PLANTS
3.1. Polyamines and Abiotic Stress
Increase in the levels of polyamines is an important characteristic of plants under different stress conditions such as salinity, chilling, drought, heat, hypoxia, heavy metals, ozone and UV [1, 2]. These changes are mainly due to changes in the biosynthesis of polyamines, oxidation of polyamines and interaction with other pathways in response to stress.
3.1.1. High-Temperature Stress
Understanding the difference between harmful and harmless changes in temperature is vital for plants to trigger protective measures against the damage. Plants are known to have molecular thermometers to sharply sense such changes [3, 4]. High temperatures lead to the accumulation of protective heat shock proteins (HSPs) to handle a stressful period of high temperature, a process known as heat shock response (HSR) [5, 6, 7]. The expression levels of polyamine biosynthetic pathway genes were studied in A. thaliana upon heat shock [8]. In the early stage of the stress, SPMS and SAMDC2 genes were found to be upregulated, which were followed by the ADC2 gene. The levels of putrescine, spermidine and spermine were increased upon heat stress, although there were no changes in the levels of thermospermine observed.
Studies on overexpressing SPMS transgenic plants and spermine deficient mutant plants revealed that higher levels of endogenous spermine are directly proportional to higher thermotolerance levels [8]. In tomato, overexpressing yeast SAMDC gene resulted in a thermotolerant plant, 55 which produced 1.7 and 2.4 fold higher levels of Spermidine and Spermine respectively, which was due to the enhancement of antioxidant enzyme activities and protection of membrane lipid peroxidation [9]. Polyamine metabolism under heat stress was also studied in tobacco plants overexpressing a proline biosynthetic enzyme, Δ1-pyrroline-5-carboxylate synthetase, which resulted in increasing the levels of putrescine, spermidine, norspermidine and spermine [10].
3.1.2. Low-Temperature Stress
In A. thaliana, responses to low temperature have been studied on gene transcription, protein, and on metabolic levels [11, 12, 13, 14, 15, 16]. In many plants, the level of polyamines increases under cold stress conditions. In wheat, alfalfa, rice, and beans, the levels of putrescine increases during the process of cold hardening [17, 18, 19]. Exogenous application of putrescine could rescue tomato cold-induced electrolyte leakage whereas the inhibitor of ornithine decarboxylase increased the leakage suggesting a protective role of putrescine in cold stress [20].
In A. thaliana, plants exposed to low temperature tend to accumulate putrescine in the first 24 hours, although no change in the levels of spermidine was observed. Related to the change in the putrescine levels are increased expressions of the polyamine biosynthetic genes ADC1 and
ADC2, after 30 minutes of cold stress, the expression of ADC1 becomes more prominent [21].
The increased levels of putrescine under cold stress are a result of increased activity of ADC1 and ADC2 [22].
3.1.3. Drought Stress
During drought conditions, the level of polyamines in plants increases resulting in a difference I the ration of different polyamines [23, 24, 25, 26, 27]. Recently, polyamines were compared to metabolites directly related to polyamine metabolism in 21 different rice cultivars 56 under long-term drought stress [27]. Under controlled conditions, putrescine dominated the polyamine pool, followed by spermidine and spermine. Under drought stress, putrescine levels decreased and spermine became the most prominent polyamine. Gene expression analysis showed that polyamine biosynthesis involving ADC responds strongly to stress when compared to the ODC pathway [27]. Overexpressing the ADC gene of Datura stramonium in rice plants improved the drought tolerance by increasing the synthesis of spermidine and spermine from putrescine [28]. In A. thaliana, a mutant deficient in producing spermine was hypersensitive to drought, which could be rescued by exogenously applying spermine and not putrescine and spermidine [29].
3.1.4. High Salinity Stress
In agriculture, soil salinity is a growing problem. Under saline conditions, the growth of plants and seed germination gets seriously affected [30]. Polyamines have been considered as important factors in salt tolerance in different plant species [31, 32, 2, 26]. In different crop species, the levels of polyamine changes under salt-stressed conditions, the levels of spermidine and spermine increases while the levels of putrescine tend to decrease. [33].
In A. thaliana, the expression levels of ADC2 and SPMS were increased under high salt conditions [34], and ADC2 is required for the accumulation of putrescine resulting in salt tolerance [35]. Exogenous application of putrescine improves in salt tolerance in rice
[36].Similar effects were seen by exogenous application of spermidine [37]. Mutant plants deficient in the biosynthesis of spermine were hypersensitive to salt stress and the effect could be reversed by application of spermine [38]. In rice, overexpressing lines that increased endogenous polyamine levels improved tolerance in high salinity conditions [39, 40]. 57
3.2. High Temperature suppresses the Delay of Flowering in PUT5:OsPUT1 plants
As mentioned in the last chapter, the phenotypes of PUT5:OsPUT1, CaMv:OsPUT1 and
CaMv:OsPUT3 showed delay in flowering for 16, 11 and 17 days respectively when grown under LDs at 22oC, when compared to the wildtype which started flowering at 27 days and put5 plants which started flowering at 23 days.
Accession Number of Rosette Leaves Time to Flowering (Days)
put5 6.01 ± .78 22 ± 2.5
Wild Type 8.95 ± 1.59 26 ± 2.5
PUT5:OsPUT1 10.9 ± 1.39 23 ± 3
CaMv:OsPUT3 11.5 ± 1.65 24± 3
CaMv:OsPUT1 10.1 ± 1.58 23 ± 4
Table 3.1. Phenotypic variation of A. thaliana genotypes at 28 .
℃ In order to understand the effect of high temperature on the delay of flowering phenotype due to the overexpression of a polyamine transporter, we grew PUT5:OsPUT1, CaMv:OsPUT1 and CaMv:OsPUT3 under LDs at 28oC along with the put5 and WT plants for comparison. We observed that under high temperature the delay of flowering phenotype disappears. Under high temperatures, the wildtype started flowering at 26 days and the put5 plants started flowering at
22 days. The promotion of flowering by the loss of PUT5 was still present at 28oC.
PUT5:OsPUT1, CaMv:OsPUT1 and CaMv:OsPUT3 flowered at 23, 23 and 24 days respectively. The number of rosette leaves was positively correlated with the time of flowering. 58
Although at higher temperature the delay in the timing of flowering disappeared, all the other phenotypic characteristics such as thicker stems, more number of leaves, more number of flowers were still present. The delay of flowering due to the higher levels of spermidine and spermine conjugates in rosette leaves is present upstream of the temperature pathway in the genetic pathways of flowering.
Figure 3.1. Increased expression of PUTs at 28 results in the disappearance of delay of flowering phenotype. A. thaliana plants at six weeks.℃
59
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CHAPTER IV.
DUAL FUNCTIONING OF PLANT ARGINASES PROVIDES A THIRD ROUTE FOR
PUTRESCINE SYNTHESIS
4.1. Abstract 65
Two biosynthetic routes are known for putrescine, an essential plant metabolite.
Ornithine decarboxylase (ODC) converts ornithine directly to putrescine, while a second route for putrescine biosynthesis utilizes arginine decarboxylase (ADC) to convert arginine to agmatine, and two additional enzymes, agmatine iminohydrolase (AIH) and N-carbamoyl putrescine aminohydrolase (NLP1) to complete this pathway. Here we show that plants can use
ADC and arginase/agmatinase (ARGAH) as a third route for putrescine synthesis.
Transformation of Arabidopsis thaliana ADC2, and any of the arginases from A. thaliana
(ARGAH1, or ARGHA2) or the soybean gene Glyma.03g028000 (GmARGAH) into a yeast strain deficient in ODC, fully complemented the mutant phenotype. In vitro assays using purified recombinant enzymes of AtADC1 and AtARGAH2 were used to show that these enzymes can function in concert to convert arginine to agmatine and putrescine. Transient expression analysis of the soybean genes (Glyma.06g007500, ADC; Glyma.03g028000 (GmARGAH) and the A. thaliana ADC2 and ARGAH genes in leaves of Nicotiana benthamiana, showed that these proteins are localized to the chloroplast. Experimental support for this pathway also comes from the fact that AtARGAH, but not AtAIH or AtNLP1, expression is co-regulated with AtADC2 in response to drought, oxidative stress, wounding, and methyl jasmonate treatments. Based on the high affinity of ARGAH2 for agmatine, its co-localization with ADC2, and typically low arginine levels in many plant tissues, we propose that these two enzymes can be major contributors to putrescine synthesis in many A. thaliana stress responses.
4.2. Introduction
A distinguishing feature of eukaryotic cells is the presence of membrane-bound organelles and vesicles. In plants, these sub-cellular spaces have multiple functions that enable the specialized compartmentation of metabolic processes such as carbon fixation, respiration, 66 energy capture, biosynthesis of primary metabolites, and the sequestering of waste products or defensive metabolites that are normally released only upon cell lysis [1-3]. One particular advantage of this organizational strategy is that the compartmentation of enzymes prevents futile cycling of metabolites by controlling the access of enzyme substrates to enzyme targets. An additional feature of plant metabolism is the redundancy of certain metabolic pathways [4-6], as is also the case for polyamine biosynthesis. In one pathway, ornithine decarboxylase (ODC) converts ornithine directly to putrescine [7] (Fig. 1). A second pathway utilizes arginine decarboxylase (ADC) to convert arginine to agmatine, and two additional enzymes, agmatine iminohydrolase (AIH) and N-carbamoyl putrescine aminohydrolase (NLP1) to synthesize putrescine [8, 9].
Putrescine serves as a precursor for the other common plant polyamines, spermidine, spermine, and thermospermine. Spermidine is required for the hypusination [10] of the three members of the elF5A family of conserved transcription elongation factors [11] that are present in all eukaryotes [12]. Overexpression of spermidine synthase in Arabidopsis thaliana increases tolerance to a wide range of abiotic stresses [13]. Thermospermine plays an essential role in normal vascular development of plants [14, 15]. Overexpression of spermine synthase results in an increased level of spermine and increased tolerance against Pseudomonas viridiflava [16].
Arabidopsis thaliana has lost ODC, and thus the second known pathway, but instead has two
ADC’s [17]. Given the multiple roles of polyamines, it is hardly surprising that homozygous mutants of both arginine decarboxylases in A. thaliana are lethal [18]. Homozygous AIH mutants are embryo defective, consistent with the hypothesis that putrescine synthesis is essential for plants [19]. However, homozygous mutants of genes encoding the terminal enzyme of the cytosolic putrescine pathway (NLP1) have no reported phenotype, and are available from the 67
Arabidopsis Biological Resource Center (ABRC). These results suggest an absolute requirement for N-carbamoylputrescine in plant metabolism, and that a second mechanism of synthesis for putrescine in A. thaliana is necessary.
In Escherichia coli and mammals, putrescine can be synthesized in a two-step process from arginine by arginine decarboxylase, which makes agmatine; and agmatinase, which converts agmatine to putrescine with the release of urea [20-22]. Enzymes with agmatinase activity are members of the ureohydrolase superfamily that also include enzymes with arginase, forminoglutamase, and proclavaminate amidoinohydrolase activities [23]. Plant arginases are unique in that they are phylogenetically more similar to bacterial agmatinases, than to bacterial or mammalian arginases [9, 24]. However, characterization of the two tomato arginase/agmatinase enzymes indicated that, at high substrate concentrations, these enzymes exhibited the highest catalytic efficiency for the conversion of arginine to ornithine and urea
[24]. Since plant arginases are known to be strongly conserved, it has been assumed that an agmatinase-dependent pathway for polyamine biosynthesis does not exist in plants.
Here we show that three arginases (ARGAH), two from A. thaliana and one from soybeans, are capable of working in concert with arginine decarboxylase to efficiently convert arginine to putrescine. This second route for putrescine synthesis in A. thaliana, and a third route for putrescine synthesis in soybeans, is localized to the plastid. In A. thaliana, ADC2 is significantly upregulated in response to drought, oxidative stress, wounding and methyl jasmonate. We also note that it is ARGAH2, and not AIH and NLP1, that increase in response to the expression of ADC2.
4.3. Materials and methods 68
4.3.1. Phyre2 Analysis
Three dimensional structure of A. thaliana ARGAH2 were predicted using Phyre2by threading the predicted protein models of ARGAH2 using the 3D template of Deinococcus radiodurans [25]. The predicted active site region of arginases/agmatinases from A. thaliana, soybeans, and poplar were aligned with active site region of D. radiodurans agmatinase using
CLUSTALW2 (http://www.ebi.ac.uk/Tools/msa/clustalw2/). Superposition of 3D structures was performed using Chimera (http://www.cgl.ucsf.edu/chimera/) [26].
4.3.2. Phylogenetic analysis
Protein sequences of ADC and agmatinase/arginase were obtained from the Phytozome database [27]. The amino acid sequence alignment was created using MUSCLE [28] .
Phylogenetic trees were constructed by MEGA 6.06 [29] using the maximum likelihood method based on the Jones-Taylor-Thornton (JTT) matrix-based model. The reliability of the trees was tested using a bootstrapping test with 1000 duplicates. Alignment files were generated using the
MULTALIN interface http://multalin.toulouse.inra.fr/multalin/multalin.html [30].
4.3.3. DNA sources and constructs
Genes were amplified using gene-specific primer pairs listed in Table S3 following the manufacturer’s guidelines for the use of Phusion® high fidelity DNA polymerase. The sequence of Glyma.03g028000 (GmARGAH) was codon-optimized for expression in yeast and synthesized by GenScript, Pistcataway, NJ. AtADC1 was amplified from genomic DNA of A. thaliana Col-0. cDNA clones of A. thaliana ADC2 (At4G34710) ARGAH1 (At4G08900) ARGAH2 (At4G08870) were obtained from the ABRC (www.arabidopsis.org). 69
4.3.4. Subcellular localization analysis
Full length sequences of AtADC2, AtARGAH2, AtADC1, GmADC, and GmARGAH, were cloned into plant expression vectors pGWB6, or pGWB5 using the GATEWAY® recombination system [31] Inserts were verified by sequencing and vectors were transformed into
Agrobacterium tumefaciens strain GV3101[32]. Samples were imaged using a Leica TCS SP5 laser scanning confocal microscope (Leica Microsystems, Bannockburn, IL) using the Leica
Application Suite Advanced Fluorescence (LASAF) program at 12-72 h after infiltration. Images were acquired in the XYZ plane in 1 µm steps with a 20X and the 63X oil objective (NA 1.40) using the sequential scan mode to eliminate any spectral overlap in the individual fluorophores.
Specifically, GFP was excited at 488 nm and detected at 510 nm. The mCherry was excited at
561 nm and detected at 610 nm. Chlorophyll autofluorescence was used to localize plastids in the cells using laser excitation at 633nm and emission of light at 670 nm. GFP signals were false- colored green and mCherry signals were false-colored red. Background fluorescence from untransformed leaves of plants at similar laser excitation settings were acquired and subtracted from images to identify fluorescence generated by tagged proteins. Images were merged using
ImageJ [33]. Images were acquired from multiple leaf sections at different time points following infiltration (24-72h) on at least two separate occasions for each construct.
4.3.5. Yeast complementation assays
The spe1 yeast knockout strain, which lacks ornithine decarboxylase (YSC6273-
201936543), was obtained from GE Darmacon, Lafayette, CO. BY4741 served as a wild-type control. Yeast strains were maintained on enriched medium (YEPD, Amresco) or minimal 70 medium (SC, Sunrise Science) supplemented with 1 mM putrescine (Sigma-Aldrich). The full- length sequence of AtADC2 was cloned into yeast expression vector pAG303GPD-ccdB
(Addgene) using the GATEWAY® recombination system. Full length sequences of AtARGAH2,
AtARGAH1 and GmARGAH were cloned into the yeast expression vector pYES-DEST52
(Invitrogen) in a similar manner. The pAG303-ADC2 plasmid construct was introduced into competent yeast spe1 mutant cells by electroporation. The resulting transformants were selected on SC lacking histidine and containing 1 mM putrescine. Positive colonies were used to produce competent cells. The pYES-DEST52-ARGAH2 plasmid construct was then introduced to the above competent cells by electroporation. The resulting transformants were again selected on SC plates lacking uracil and containing 1 mM putrescine. spe1 yeast mutant cells containing pAG303-ADC2 and pYES-DEST52-ARGAH, pAG303-ADC2 and pYES-DEST52-
GmARGAH were prepared and tested in a similar manner. Cultures of wildtype (WT), spe1, and spe1 transformed with ADC2 alone, spe1∆ transformed with ARGAH1, ARGAH2, or
GmARGAH; or spe1 transformed with both ADC2 and one of the arginases were first grown overnight on YPD plates. To deplete cells of polyamine stores, single colonies from the YPD plates were grown overnight in 5 ml of SC media with 1% raffinose and 2% galactose without polyamines. A dilution of 1 x 10-4 of the cells was prepared in the same medium. Aliquots of each dilution were then streaked onto SC plates with 1% raffinose, 2% galactose, and SC plates with 1% raffinose, 2% galactose, and 1 mM putrescine. The plates were incubated overnight at
30 ºC and photographed after 24 hours.
4.3.6. Growth rate of yeast strains without polyamine supplementation
The wild type yeast strain BY4741, spe1yeast knockout strain and spe1 yeast mutant containing pAG303-AtADC2 and pYES-DEST52-AtARGAH2 were maintained on YPD 71 medium. A single colony from each strain was transferred into liquid SC minimal medium with
2% galactose. The cultures were allowed to grow overnight at 30 ºC. One ml of a 10-4 dilution from each culture was then used in a BioTek Synergy HT plate reader to measure the absorbance of the cultures at 600 nm every one hour over a period of 18 hours. Standard curves were generated using the absorbance data, and used to estimate the doubling time of each strain.
4.3.7. Agmatinase activity assays
AtARGAH2 was cloned into pBAD-DEST49 Gateway destination vector according to the manufacturer’s instructions. The enzyme was purified using HIS-Select™ kit from Sigma-
Aldrich, as described in the manual. Enzyme assays were performed at 25°C in 10 mM Tris-HCl, pH 8.0, 1mM MnCl2 in order to reflect ambient conditions in the chloroplast during the light period. Dansyl derivatization [34] of the enzyme assay samples were performed as follows. A
200 µl aliquot of the reaction mix was mixed with 40 µl of 0.05 mM 1,7-heptanediamine
(Sigma-Aldrich) as an internal standard, 200 µl of saturated sodium carbonate (13%, w/v) and
400 µl of dansyl chloride in acetone (10 mg/ml). After vortexing, the mixture was incubated overnight at room temperature in the dark. Excess dansyl chloride was removed by adding 100
µl of L-proline (100 mg/ml; Sigma-Aldrich) followed by an incubation for 30 min at room temperature in the dark. The dansylated amines were further extracted with 500 µl of toluene.
The extraction was repeated two times. The organic phase containing amines was vacuum evaporated and the residue was dissolved in 500 µl of methanol. The dansylated amines were separated and quantified by HPLC with an Agilent Technologies 1120 series HPLC. A Gemini®
5 µm C18 110 Å, 250 × 4.6 mm LC column was used for the separation of polyamines. Samples were eluted from the column with a programmed water: methanol solvent gradient over 30 minutes. The initial conditions were 10% methanol and 90% water pumping at a flow rate of 72
0.75 ml min-1. The methanol concentration was increased to 60% over 4 minutes and then up to
80% over 11 minutes. After that, the concentration of methanol was further increased to 95% over 10 minutes. These conditions were kept constant for 1 minute and then returned to initial conditions. After each cycle, the column was washed with 100% methanol for five minutes and re-equilibrated for five minutes. A standard curve was generated by measuring known amounts of putrescine and used for the estimation of putrescine content. Agmatinase activity was determined by measuring the time-dependent conversion of 250 µM agmatine to putrescine by
HPLC, after correction for baseline contamination of putrescine in the agmatine sample. Kinetics of putrescine production by ARGAH2 was estimated over the substrate range of 10-250 µM agmatine. Michaelis–Menten parameters were calculated using a non-linear regression method with GraphPad Prism software (http://www.graphpad.com/prism/Prism.htm).
4.3.8. In vitro assay of ADC2 and ARGAH2
AtADC2 and AtARGAH2 were cloned separately into pBAD-DEST49 Gateway destination vector according to the manufacturer’s instructions. The enzymes were purified using
HIS-Select™ kit from Sigma-Aldrich, as described in the manual. Enzyme assays were performed at 25°C in a buffer modified from [24]. Two hundred µl of the assay buffer containing
10 mM Tris, 1 mM MnCl2 and 1 mM arginine (pH 8) was mixed with 100 µl of arginine decarboxylase and 100 µl of arginase/agmatinase. The reaction mixture was incubated at 25 ºC for 45 minutes. After incubation, 400 µl of 5% perchloric acid was added to stop the reaction.
Dansyl derivatization [34] of the enzyme assay samples were performed as described above.
Samples were eluted from the column with a programmed water: methanol solvent gradient over
30 minutes. The initial conditions were 10% methanol and 90% water pumping at a flow rate of
0.75 ml min-1. The methanol concentration was increased to 70% over 4 minutes and then up to 73
85% over 10 minutes. These conditions were kept constant for 1 minute and then the concentration of methanol was further increased to 95% over 8 minutes. Finally, the concentration was returned to the initial conditions. After each cycle, the column was washed with 100% methanol for five minutes and re-equilibrated for five minutes. Data collection was started after the arginine peak was eluted. This program enabled baseline separation of agmatine, ornithine and putrescine.
4.4. Results
4.4.1. Plant arginases have agmatinase activity
It was previously noted that plant arginases cluster phylogenetically with agmatinases, rather than arginases from other Kingdoms [24]. We noted that BLAST analysis of the A. thaliana ARGAH2 protein has best hits to predicted bacterial agmatinases with sequence identities in the range of 43-47% and sequence similarities in the range of 63-67%. We next used the Protein Recognition server PHYRE2 [25] with plant arginases from the Phytozome database
[27] to identify a bacterial sequence from D. radiodurans for which there is also a crystal structure [23]. Sequence alignment of arginases from A. thaliana, Glycine max (soybean), and
Populus trichocarpa, with Deinococcus radiodurans showed a high level conservation of amino acid sequences, including those known to be critical for enzymatic activity (Fig. 2). We used
PHYRE2 to build 3D models of ARGAH2 from A. thaliana, and D. radiodurans [25].
Superimposition of the Arabidopsis ARGAH2 with the 3D structure of D. radiodurans
(DR) agmatinase [23] showed high structural similarity (Fig. 3, Supplementary Movie 1). In prior work, the conservation of critical amino acids in the human agmatinase relative to those of
D. radiodurans had been noted [21]. In a similar manner, key residues of the ARGAH2 protein 74
were well superimposed with those of DR agmatinase: His163 (His121 in DR agmatinase); Asp187
(Asp143); Pro190 (Leu146); Asp191 (Asp147); Tyr193 (Thr149); His203 (Asn159); Asn228 (Asp187);
Asp272 (Asp227); Asp274 (Asp229); His286 (Ser243); and Glu315 (Glu274) (Fig. 3) [23]. Conservation of these residues in the predictive active site of this proteins was also observed in soybean, tomato, poplar, and rice (Fig. 2). Based on this evidence, we hypothesized that these plant arginases should have agmatinase activity.
Co-transformation of an oat ADC and either a human or an E. coli agmatinase gene has previously been used to reconstitute a putrescine biosynthetic pathway in yeast [21]. We adopted a similar strategy to assess the potential agmatinase activity of the arginases AtARGAH2
(At4g08870), AtARGAH1 (At4g08900), and GmARGAH, (Glyma.03G028000). The yeast ODC mutant, spe1, does not grow in the absence of exogenous putrescine once polyamine stores have been depleted [35]. However, co-expression of each of AtARGAH1, AtARGAH2, and GmARGAH with AtADC2, fully complemented the spe1growth defect of (Fig. 4). Thus, both arginases from
A. thaliana and at least one arginase from soybean are capable of synthesizing putrescine when supplied with agmatine. To evaluate the efficiency of these yeast constructs to support rapid growth in continuous culture, we measured the growth rate of WT, spe1, and spe1 strains expressing the plant genes AtADC2 and the three ARGAH genes after transfer to minimal medium. With depletion of putrescine, the spe1 strain ceased growth (Fig. 5). In the absence of exogenous putrescine, the doubling time of the WT strain was 3.910.35h, while the yeast strains expressing ADC2 and ARGAH1 or ARGAH2 had a doubling time of 4.7 ± 0.35h and 4.4 ± 0.3h respectively. The doubling time for yeast strains expressing ADC2 and GmARGAH was 5.02 ±
0.25h. We also noted that the growth of the transformed spe1 yeast strains plateaued at a lower cell density than WT cells, which may indicate the accumulation of a toxic intermediate. Taken 75 together, the plate complementation and growth assays show that these plant genes form a viable metabolic pathway for putrescine synthesis in yeast cells.
The agmatinase activity of AtARGAH2 at low substrate concentrations was confirmed by
HPLC analysis of the products. The affinity-purified enzyme converted 250 µM agmatine to putrescine in a time-dependent manner (Fig. S1a). The enzyme had a high affinity for agmatine
(Fig. S1b) with a Km estimated at 72.6 ±10 µM. The affinity of ARGAH2 for agmatine is considerably lower than the affinity of the tomato arginases for arginine (Km = 29-32 mM) [24].
Since plant arginases have a high level of sequence conservation (Fig. 2), all plant arginases are likely to have a similar affinity for the substrates arginine and agmatine.
Soybean arginine decarboxylases have a much higher affinity for arginine (Km = 46.1
µM) [36], than that reported for arginases suggesting that even when ADC and ARGAH are localized together, arginine would be preferentially metabolized by ADC at low concentrations of arginine. To confirm that ADC2 and ARGAH2 could form a biosynthetic route from arginine to putrescine without significant accumulation of ornithine, we affinity-purified both proteins and added them together in the presence of 1 mM arginine. A chromatographic method was developed to enable HPLC separation of agmatine, ornithine, and putrescine (Fig. 6A). HPLC analysis of the dansylated substrates and products confirmed the production of agmatine by
ADC, and putrescine by ARGAH, with a very small peak of ornithine, indicating a low level of arginase activity (Fig. 6B).
4.4.2. ADC2 and ARGAH2 together form a plastid putrescine pathway
We used GFP fusion constructs of ADC2 and ARGAH2 to determine whether these enzymes are co-localized in the cell. Transient expression in Nicotiana benthamiana leaves of 76 these GFP fusions, showed that the proteins were localized to the chloroplasts (Fig. 7). Thus, the supply of putrescine synthesized by this pathway is controlled by transport from the chloroplast.
4.4.3. The plastid pathway is activated in stress responses
The two ADCs of A. thaliana also show a strong divergence in expression [37-40].
However, agmatine produced by ADC2 in the plastid could be converted to putrescine by agmatine in the chloroplast, or exported to the cytoplasm for conversion to putrescine by AIH and NLP1. To evaluate which biosynthetic strategy is preferred in A. thaliana, we used E-
Northern analysis to determine the likely contributions of putrescine synthesis of each pathway
[41]. Under four conditions when ADC2 is strongly upregulated, drought, oxidative stress, wounding, and application of methyl jasmonate, increased ADC2 expression was correlated with increased ARGAH2 expression (Fig. S3). In contrast, expression of AIH which converts agmatine to N-carbamoylputrescine, stayed the same or declined. Notably, we were unable to identify conditions where the increased expression of ADC1 or ADC2 was correlated with increased expression of AIH and NLP1 (not shown). Thus, in summary, the likely existence of a functional plastid pathway for polyamine biosynthesis in A. thaliana is supported by co-localization of the necessary enzymes, complementation experiments in yeast, kinetic analysis demonstrating the high affinity that ARGAH2 has for agmatine; and four examples showing co-expression of
ARGAH2 and ADC2 in A. thaliana in response to various stresses.
4.4.4. Soybeans also have a complete plastid putrescine pathway, but most plants do not
To determine whether a complete biosynthetic pathway for putrescine in the chloroplast might exist outside of the Brassicaceae, we used transient expression assays of GmADC2
(Glyma.06G007500) and GmARGAH (Glyma.03G028000) from soybean in N. benthamiana 77 leaves to show that these genes are also localized to the chloroplast (Fig. 7). Thus soybeans, along with A. thaliana have a complete chloroplast-localized putrescine biosynthetic pathway.
To further investigate the role of this pathway in putrescine biosynthesis for other plants, we used the Phytozome database [27] to identify species with multiple ADC and ARGAH genes.
We found that, 23 of 43 sequenced plant genomes have more than one ADC, and 12 of these genomes have at least one ADC with predicted localization to the plastid (Table S1). Within this subset of plant genomes, 13 genomes include more than one gene with predicted ARGAH activity. Phylogenetic analysis shows that all of the ADCs from seven brassica genomes form two clades of nine and five sequences (Fig. S3). AtADC2 is in a clade separate from AtADC1, and all the members of the clade with AtADC2, which include at least one gene from each species, are predicted to be localized to the plastid. Other dicot species with two ADC genes include cacao, strawberry, tomato, potato, soybeans and Mimulus guttatus. With the exception of soybeans, at least one of the ADCs in each of these species, has a predicted plastid localization.
The monocots; Zea mays, Oryza sativa, and Panicum virgatum also contain two ADC genes, but none of these genes are predicted to be targeted to the chloroplast. However in oats, immunological staining indicated that ADC was localized in the plastid [42], so other monocots may have chloroplast-localized ADCs. We also noted that Physcomitrella patens has three predicted ADC genes, and one of them is predicted to be localized in the chloroplast. A smaller number of species were found to have two genes with predicted ARGAH activity (Fig. S4).
Species that contain two members of both the ADC and ARGAH gene families included four members of the Brassicaceae, along with tomatoes, and soybeans. Dicots that have duplications in only ARGAH, but not ADC, include cassava, flax, and poplar. Each of these species includes at least one ARGAH with a predicted localization to the chloroplast (Table S2). In these species, 78 putrescine can be synthesized in the chloroplast, if agmatine is transported into the chloroplast, or if the single ADC is localized there. Notably, no monocot species was identified with more than one ARGAH. Thus the retention of a complete plastid pathway is relatively uncommon in plant genomes. Of the seven sequenced Brassica genomes with two ADC’s, only four were found to have retained two ARGAH genes.
4.5. Discussion
All plant ARGAH enzymes are very strongly conserved throughout the length of the protein sequence (Fig. S3) and [24]. We suggest that the need for this level of sequence conservation is due to the dual specificity of these enzymes. ARGAHs can function as arginases but this activity only occurs at high (mM) levels of arginine [24]. The two ARGAH enzymes of
Lycopersicon esculentum (tomato) were characterized as arginases with a pH optimum of 9.5 and
Km of 32 ± 4 and 29 ± 6 mM for LeARG1 and LeARG2, respectively. At 100 mM substrate concentrations of arginine or agmatine, these enzymes preferentially metabolized arginine to ornithine and had very low relative levels of agmatinase activity. Given the high level of sequence conservation of plant arginases, (Fig. 2) it is likely that all plant enzymes have a similarly high Km for arginine and function as arginases when arginine is available at high levels.
Arginine is a major storage amino acid in seed storage tissues such as soybeans [43], and peas, where it constituted 66% of the total amino acid pool [44]. The arginase activity of this enzyme enables germinating seedlings to catabolize arginine that has accumulated in the developing seeds [45, 46]. On the other hand, arginine levels in plant leaves are a small fraction of the total pool of amino acids and are thus not high enough to support significant levels of arginase activity[47-49]. Phloem transport to developing tissues is also not a significant source of 79 arginine [49, 50]. In most tissues, arginine levels are often less than 1 mM, and the kinetic properties of plant arginases [24] indicate that, at 1 mM arginine, arginase activity would only be about 3% of the maximal rate . Agmatine levels have been determined in eight brassica species
[51]. For example, in broccoli, leaf agmatine levels were in the range of 0.2-0.35 µmoles/g wet weight. Lower levels of agmatine were consistently reported in infected tissues, presumably reflecting its conversion to putrescine and other polyamines. The estimated Km for ARGAH2 reported here (72.63 ±10 µM) and that of agmatine iminohydrolase (AI, Km = 112 µM) [8] suggests that both ARGAH and AIH are capable of metabolizing agmatine at substrate levels found in these tissues. We suggest that the very high level of sequence conservation for this enzyme across the plant kingdom is unlikely to be due to selection pressure against mutations that would decrease its arginase activity, but rather the requirement for the enzyme to retain activity for both arginine and agmatine.
The yeast complementation assays described in this paper, mirror earlier experiments demonstrating the agmatinase activity of the human enzyme [21]. Moreover, the plant enzymes supported a significant growth rate in yeast. HPLC analyses of ARGAH2 alone, and ARGAH in combination with ADC2 confirm the ability of ARGAH to convert agmatine to putrescine at low substrate concentrations (Fig. 6a). Metabolic analyses of ADC and ARGAH mixtures in these in vitro assays also indicates that they don’t directly compete for arginine, since we detected only a small ornithine peak, but enable the synthesis of putrescine in a two-step process (Fig. 6b). In the presence of 1 mM arginine, we did not observe any significant accumulation of ornithine in the in vitro assays of the two enzymes (Fig. 6b). Thus, at low substrate concentrations of arginine and agmatine, ARGAH acts preferentially as an agmatinase. In tomatoes, LeARG2, but not
LeARG1 is strongly induced in response to wounding, methyl jasmonate and the phytotoxin 80 corontine [24]. Coronotine is a functional analogue of jasmonate-isoleucine conjugate, and thus the induction of LeARG2 in response to wounding and methyl jasmonate follows the same pattern of expression that is seen for AtARGAH2 (Fig. S2). The co-expression of ARGAH with
ADC by coronatine likely enables the direct synthesis of putrescine and not ornithine, since arginine levels in source leaves are typically too low to support significant levels of arginase activity.
Increased synthesis of putrescine contributes to protection of fruit from chilling stress
[52]. The up-regulation of LeARG1 and LeARG2 in tomato fruits that occurs in response to chilling [53] likely contributes to the synthesis of putrescine rather than ornithine, as soluble levels of arginine in ripe tomatoes are also less than 1 mM [54]. A consequence of chilling stress is the increased production of reactive oxygen species (ROS). Polyamines are a source of H2O2 via the metabolism of polyamine oxidases [55]. Elevated levels of polyamines can also mitigate the effects of ROS via multiple mechanisms [56]. In A. thaliana expression of ADC2 and
ARGAH2, but not ADC1, is strongly induced by oxidative stress (Fig. 9). To summarize, the up- regulation of ARGAH in tissues with low levels of arginine, and its co-expression with ADC certainly hints that this may be a major pathway for putrescine synthesis in response to certain stresses.
Analysis of A. thaliana ADC mutants does not directly support the physiological contribution to putrescine synthesis by agmatinase. However, the fact that the two ADCs are differentially expressed, and that ADC2 is compartmentalized with ARGAH2, is exactly what is expected if the activity of ADC2 is intended to support a second pathway. Notably, adc1 and adc2 mutants show no adverse phenotypes in the absence of abiotic challenges, but both mutants are more sensitive to freezing [39]. In response to cold, both AtADC1 and AtADC2 are up- 81 regulated, but the plastid-localized enzyme AtADC2 is activated to a lesser extent, and appears to be activated by abscisic acid (ABA)-responsive elements. adc2 mutants show increased sensitivity to several kinds of abiotic stresses. These include, reduced accumulation of putrescine in response to high levels of NaCl, and increased sensitivity to salt stress [57] [37]. Knockout mutations of adc2 resulted in a 20-fold decrease in ADC activity in response to osmotic stress
[38]. This indicates that ADC2 is the major contributor to putrescine synthesis in response to osmotic stress. Since whole plant extracts from Ds insertion mutants of ADC2 had levels of putrescine that were only 25% of wild type, this gene is a major contributor to steady state polyamine biosynthesis [37].
Insights on the role of the chloroplast putrescine biosynthesis pathway in A. thaliana can also be gleaned from expression analysis of ARGAH. GUS staining revealed high levels of expression of both ARGAH genes in developing leaves of Arabidopsis and especially in the vascular tissues [58] [59]. We hypothesize that the high level of expression of ARGAH genes in developing tissues reflects their role in putrescine synthesis rather than arginine catabolism, since arginine levels in these tissues are not high [48, 50]. Elevated expression of ARGAH2 was also seen in response to methyl jasmonate [59] a treatment that also results in up-regulation of ADC2
(Fig. 9). ARGAH2 expression was strongly induced following the inoculation of leaves with
Botrytis cinerea [60], and transcript levels of ARGAH1 and ARGAH2 are significantly up- regulated in response to ABA, drought stress, cold, and NaCl [61]. These are also conditions that result in the induction of one or more of the ADCs [62]. The contribution of ARGAH2 to polyamine levels and disease resistance has also been assessed using both silenced and overexpression lines [60]. However, both overexpressed lines and silenced lines, were identified with significantly higher levels of putrescine. This could be a consequence of compensatory 82 mechanisms, where down-regulation of putrescine synthesis by one pathway results in up- regulation of the other pathway. Notably, two overexpression lines showed increased resistance to Botrytis cinerea. Both overexpressed and silenced lines were also identified with lower levels of arginine and ornithine, and higher levels of proline. These results indicate that other factors in addition to ARGAH regulate the levels of these amino acids. Both susceptible and partially resistant accessions of A. thaliana to the obligate pathogen Plasmodiophora brassicae, showed a significant up-regulation of the expression of ADC1,ADC2, AIH, NLP1, SPDs1, SPDS2, SPMS as well as ARGHA1and ARGAH2 [63]. However, at 21 days after infection, arginase activity was more than 10-fold higher in susceptible roots than in partially resistant roots, and agmatine levels in susceptible roots were lower than in partially resistant roots. In contrast, arginase activity did not affect arginine levels of root tissues. The induction of arginase was interpreted as a pathogenic strategy of P. brassicae, to alter arginine metabolism, but at the time, the role of this enzyme in the conversion of agmatine to putrescine was not known. We have noted that the genome sequence of this pathogen [64] contains three genes with homology to polyamine uptake transporters that are also found in plants, stramenopiles and alveolates [65]. Thus, the induction of arginase activity in the plant may be a means of supplying putrescine to this obligate pathogen.
All sequenced plant genomes have a cytosolic pathway for putrescine synthesis, but the brassicas have lost ODC. The presence of a cytosolic pathway in A. thaliana may indicate a particular requirement for the synthesis of specific putrescine conjugates, since the N- carbamoylputrescine intermediate in this pathway (Fig. 1) is a more reactive compound for conjugation, than putrescine itself, and thus may be the actual substrate used in the formation of some putrescine conjugates. Notably, there are embryo-deficient mutants of AIH [19, 66] but no 83 known phenotype is associated with homozygous mutants of NLP1, the terminal step in the cytosolic putrescine biosynthesis pathway. Arabidopsis thaliana plants with mutations in both
ARGAH genes are still viable, indicating that synthesis of putrescine by these enzymes is dispensable under some conditions [61] and that, under certain conditions, the loss of putrescine synthesis from one pathway can be partially compensated by an alternate pathway for putrescine synthesis in A. thaliana. However in rice, which has only a single gene encoding arginase/agmatinase activity, loss of OsARG function resulted in plants with reduced height, smaller panicles, and smaller grain size [67]. Conversely, the authors also reported that overexpression of arginase increased seed set and grain filling of field-grown plants under N- limited conditions. The highest expression of OsARG was found in the panicles, where OsARG is hypothesized to function in arginine recycling of nitrogen that is transported to the phloem from other parts of the plant. In the null mutants of this gene, arginine levels, which represented only 1% of the total amino acid pool in the panicles of WT plants, rose more than 9.7 fold. This accumulation of arginine in the mutants was attributed to the loss of arginase activity. However other arginine-utilizing pathways may also be affected by the loss of active OsARG. Expression analyses of the two rice ADC genes [41] indicate that both are up-regulated during panicle development. Because OsARG, like all plant arginases, is also capable of converting agmatine to putrescine, the loss of agmatinase activity, could result in feedback inhibition of ADC due to the accumulation of agmatine [36].
Several dicot genomes contain more than one arginase (Table S2), although there is no correspondence between those genomes and genomes with multiple ADCs. Notably, there are no sequenced monocot genomes in the Phytozome database [27] with more than one agmatinase gene. Perhaps this reflects a particular requirement for monocots to control where in the cell 84 putrescine is synthesized. Given the frequency of interspecies hybridization events, and whole genome duplications in the evolutionary history of plants [68] it is noteworthy that while approximately half of the sequenced genomes in the Phytozome database contain two ADCs, and one of the ADC’s has a predicted plastid location (Table S1) many species do not have two copies of both enzymes.
Here we noted that transient expression of ARGAH2 and GmARGAH in N. benthamiana leaves showed that these enzymes are localized to the chloroplast in leaf tissues. The chloroplast localization of this enzymatic activity has also been noted in two proteomic studies [3, 69].
However, ARGAH1 and ARGAH2 were also localized to the mitochondria of cultured tobacco cells [58]. A large number of proteins that are targeted to both the mitochondria and the plastid
[70, 71] have now been identified. Since all plant ARGAH enzymes have agmatinase activity, arginases that are localized to the mitochondria may also provide putrescine for that organelle, if agmatine can be transported in.
The presence of two differentially regulated, and spatially separated, arginine decarboxylases in A. thaliana could provide a mechanism for the synthesis of two different putrescine pools. Delivery of agmatine to the mitochondria could also provide a direct source of putrescine for that organelle. Since the relative affinity for agmatine of AIH and ARGAH is in the same range, the importance of these enzymes in the synthesis of putrescine under different conditions needs to be addressed by metabolic flux analyses.
4.6. Acknowledgements 85
We thank Arabidopsis Biological Resource Center for providing seeds and clones, and
Gerald Jones for assistance with HPLC analysis. We also thank Georg Jander for many insightful comments. This work was supported by The Graduate College, BGSU.
4.7. Accepted Manuscript
Title: Dual functioning of plant arginases provides a third route for putrescine synthesis
Authors: Jigar Patel, Menaka Ariyaratne, Sheaza Ahmed,
Lingxiao Ge, Vipaporn Phuntumart, Andrea Kalinoski, Paul F.
Morris
PII: S0168-9452(17)30049-3
DOI: http://dx.doi.org/doi:10.1016/j.plantsci.2017.05.011
Reference: PSL 9609
To appear in: Plant Science
Received date: 24-1-2017
Revised date: 4-5-2017
Accepted date: 25-5-2017
Please cite this article as: Jigar Patel, Menaka Ariyaratne, Sheaza Ahmed, Lingxiao Ge,
Vipaporn Phuntumart, Andrea Kalinoski, Paul F.Morris, Dual functioning of plant arginases 86 provides a third route for putrescine synthesis, Plant Science http://dx.doi.org/10.1016/j.plantsci.2017.05.011
87
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Figure 4.1. Biosynthetic strategies for the synthesis of putrescine.
Figure 4.2. Plant agmatinases show conservation of key active site residues with agmatinase from Deinococcus radiodurans. 97
Figure 4.3. Predicted 3D structure of AtARGAH2 is highly similar to the crystal structure of Deinococcus radiodurans (DR) agmatinase. (A) Crystal structure of DR agmatinase (B)
Predicted 3D structure of A. thaliana (AT) agmatinase (C) Superimposition of DR and AT agmatinase (D) Active site residues of DR agmatinase (E) Predicted active site residues of AT agmatinase (F) Superimposition of DR and AT agmatinase active site residues. Structure of
AtARGAH2 was generated by threading over DR agmatinase using PHYRE2 [25].
Superposition of DR agmatinase and AT agmatinase was done with the UCSF Chimera package
[26].
98
Figure 4.4. Characterization of plant agmatinases by complementation in yeast. Strains were plated on SC minimal media in the presence and absence of exogenous putrescine.
(A) BY4741, spe1, spe1+ AtADC2, spe1 +AtARGAH2 and spe1 + AtADC2 +AtARGAH2.
(B) BY4741, spe1, spe1+ AtADC2, spe1 +AtARGA12 and spe1 + AtADC2 +AtARGA1.
(C) BY4741, spe1, spe1+ AtADC2, spe1 +GmARGAH and spe1 + AtADC2 +GmARGAH. 99
Figure 4.5. Growth of yeast strains in the absence of putrescine. Indicates that expression of
ADC2 and ARGAH2 in spe1 mutants supports rapid growth in liquid medium.
Figure 4.6. A. HPLC analysis confirms agmatinase activity of AtARGAH2. Affinity-purified
AtARGAH2 shows time-dependent conversion of agmatine to putrescine. B. in vitro assay of agmatinase activity of ARGAH2. The apparent Km value for agmatine is 74.8 ±µM.
100
.
Figure 4.7. HPLC analysis of arginine decarboxylase and arginase/agmatinase show accumulation of agmatine and putrescine. (A). HPLC separation of standards: agmatine 7.2 min, ornithine 8.1 min; putrescine 10.1 min; internal standard,1,7 diaminoheptane,13.9 min. (B)
HPLC profile of metabolites produced by arginine decarboxylase and ARGAH2 in the presence of 1 mM arginine. Data collection was started after the elution of the arginine peak. In (A) the concentrations are agmatine 10 µM, putrescine 10 µM, ornithine 10 µM and internal standard 4
µM. In (B) the estimated concentration of the products are: agmatine 7 µM, putrescine 3 µM, using an internal standard of 5 µM. 101
Figure 4.8. Confocal microscopy showing plastid localization of AtADC2 and AtARGAH2 in transiently expressed N. benthamiana leaves. (A) Chlorophyll autofluorescence (B) ADC2-
GFP (C) Brightfield, (D) Merged image, (E) Chlorophyll autofluorescence, (F) AtARGAH2-
GFP, (G) Bright field, (H) Merged image.
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Figure 4.9. In four stress responses of A. thaliana, ARGAH2 expression clusters with
ADC2. E-northern heat map analysis of log transformed and clustered genes associated with putrescine biosynthesis from BAR [37] . Increased expression of genes in the heat map is indicated by a gradient of orange to red, while down-regulation of gene expression is indicated by darker shadings of grey. The heat map data in each experiment is optimized to emphasize the difference in expression for each data set. A. Drought response of shoots B. Oxidative stress response of shoots. C. Wounding response of shoots. D. Response of whole plants to methyl jasmonate.
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Figure 4.10. Confocal microscopy showing chloroplast localization of GmADC2 and
GmARGAH in transiently expressing N. benthamiana leaves. (A) GmARGAH-GFP (B)
Chlorophyll autofluorescence (C) Merged image (D) GmADC2-GFP (E) Chlorophyll autofluorescence and (F) Merged image.
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Figure 4.S1. Maximum likelihood phylogeny of selected plant arginine decarboxylases.
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Figure 4.S2. Maximum likelihood phylogeny of selected plant arginase/agmatinases.
106
107
108
Figure 4.S3. Fasta alignment file of plant species with Arginine decarboxylases. Species designations for individual genes are as follows: Physcomitrella patens, Phpat.006G071900,
Phpat.005G013600, Phpat.016G006200; Oryza sativa LOC_Os06g04070, LOC_Os04g01690;
Panicum virgatum, Pavir.J13691, Pavir.J33899; Zea mays GRMZM2G396553,
GRMZM2G374302; Mimulus guttatus Migut.N01279, Migut.B00171; Solanum lycopersicum
Solyc10g054440.1, Solanum lycopersicum Solyc01g110440.2; Solanum tuberosum
PGSC0003DMG400026671, Solanum tuberosum PGSC0003DMG400001662; Theobroma cacao Thecc1EG006773, Thecc1EG020310; Arabidopsis lyrata 491183, 931833; Arabidopsis thaliana, AT4G34710 (ADC2), AT2G16500 (ADC1); Brassica napa BnaA03g53010D,
BnaC07g45200D; Brassica rapa, Brara.K00332, Brara.A00352; Capsella grandiflora,
Cagra.2350s0020, Cagra.21579s0001; Capsella rubella Carubv10004246m.g,
Carubv10013090m.g; Eutrema salsugineum,Thhalv10024529m.g; Eutrema salsugineum
Thhalv10022578m.g; Fragaria vesca gene00390-v1, gene01668-v1.0-hybrid; Glycine max
Glyma.06G007500; Glyma.04G007700.
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Figure 4.S4. Fasta alignment file of plant arginase/agmatinases. Aligned by MultAlign.
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Species Name Gene PLPred TargetP Predota WolfPSo Arabidopsis lyrata 491183 Plastid Plastid Arabidopsis lyrata 931833 Plastid Plastid Arabidopsis thaliana AT4G34710 Plastid Arabidopsis thaliana AT2G16500 Plastid Plastid Plastid Brassica napa BnaA03g53010D Plastid Plastid Plastid Brassica napa BnaC07g45200D Plastid Plastid Plastid Brassica rapa Brara.K00332 Plastid Plastid Plastid Brassica rapa Brara.A00352 Plastid Capsella grandiflora Cagra.2350s0020 Plastid Plastid Plastid Capsella grandiflora Cagra.21579s0001 Plastid Capsella rubella Carubv10004246m.g Plastid Plastid Plastid Capsella rubella Carubv10013090m.g Plastid Eutrema salsugineum Thhalv10024529m.g Plastid Plastid Plastid Eutrema salsugineum Thhalv10022578m.g Plastid Fragaria vesca gene00390-v1.0-hybrid Plastid Plastid Fragaria vesca gene01668-v1.0-hybrid Plastid Plastid Glycine max Glyma.06G007500 Plastid Glycine max Glyma.04G007700 Plastid Mimulus guttatus Migut.N01279 Plastid Plastid Mimulus guttatus Migut.B00171 Plastid Oryza sativa LOC_Os06g04070 Oryza sativa LOC_Os04g01690 Panicum virgatum Pavir.J13691 Plastid Panicum virgatum Pavir.J33899 Physcomitrella patens Phpat.006G071900 Physcomitrella patens Phpat.005G013600 Physcomitrella patens Phpat.016G006200 Plastid Solanum lycopersicum Solyc10g054440.1 Plastid Solanum lycopersicum Solyc01g110440.2 Plastid Solanum tuberosum PGSC0003DMG400026 Plastid Plastid Solanum tuberosum PGSC0003DMG400001 Plastid Plastid Theobroma cacao Thecc1EG006773 Plastid Plastid Plastid Theobroma cacao Thecc1EG020310 Plastid Plastid Zea mays GRMZM2G396553 Zea mays GRMZM2G374302 Plastid
Table 4.S1. Predicted plastid localization of arginine decarboxylase genes from sequenced plant genomes by PLPred, TargetP, Predotar and WolfPsort.
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Species Name Gene PLPred TargetP Predotar WolfPSort Arabidopsis lyrata 489723 Plastid Plastid Arabidopsis lyrata 489781 Plastid Arabidopsis thaliana AT4G08900 Plastid Arabidopsis thaliana AT4G08870 Plastid Brassica napa BnaC03g28300D Plastid Plastid Brassica napa BnaA03g23800D Plastid Plastid Capsella rubella Carubv10001349m.g Plastid Capsella rubella Carubv10001337m.g Plastid Glycine max Glyma.17G131300 Plastid Glycine max Glyma.01G140200 Plastid Glycine max Glyma.03G028000 Plastid Linum usitatissimum Lus10000795.g Plastid Linum usitatissimum Lus10030288.g Plastid Plastid Manihot esculenta cassava4.1_011300m.g Plastid Manihot esculenta cassava4.1_011323m.g Plastid Plastid Populus trichocarpa Potri.002G146200 Plastid Plastid Populus trichocarpa Potri.014G067700 Plastid Solanum lycopersicum Solyc01g091160.2 Plastid Plastid Solanum lycopersicum Solyc01g091170.2 Plastid Table 4.S2. Predicted plastid localization of arginase/agmatinase genes from sequenced
plant genomes by PLPred, TargetP, Predotar and WolfPsort.
PCR targets Forward Reverse
AtARGAH1 CACCATGTCGAGGATTATTGGTAGAAAAGG TTTCGAGATTTTCGCAGCTAAT
AtARGAH2 CACCATGTGGAAGATTGGGCAGAG TTTTGACATTTTTGCGGCTAGC
AtADC2 CACCATGCCTGCTTTAGCTTGC TCACGCAGAGATGTAATCGTAG
GmARGAH CACCATGAGTTTCCTTCGTTCTTT TTTTGACATCTTTGCAGCGAGTTCT
GmODC CACCATGCCTTCACTAGTTG TTAGAACATGCTGTGTTCCG
OsODC CACCATGGCAGCAGCAGAA GGTGGCACTTTCCCAGTGAT
Table 4.S3. Primers used to amplify gene targets.
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CHAPTER V.
ATPDR11 IS A BROAD SUBSTRATE TRANSPORTER OF POLYAMINES AND
SELECTED AMINO ACIDS
5.1. Introduction
ABC transporters are found in all organisms and are considered to be a big family including 79 members in Escherichia coli, 29 in Saccharomyces cerevisiae, and 131 in
Arabidopsis thaliana. The transporter uses ATP hydrolysis to transport various substrates across membranes such as including lipids, heavy metal ions, inorganic acids, glutathione conjugates, sugars, amino acids, peptides, secondary metabolites, and xenomolecules, used as drugs [1, 2].
Based on the phylogenetic analysis and structure, ABC genes are divided into different subfamilies. The names of the subfamilies are often related to the function of the gene like drug transport, channel or channel regulator. The three best characterized subfamilies of the ABC gene families are multidrug resistance (MDR), multidrug resistance-associated protein (MRP) and the pleiotropic drug resistance (PDR) subfamilies.
ABC proteins have a characteristic modular structure consisting of a single or double set of two basic structural elements: a hydrophobic transmembrane domain usually made up of six membrane-spanning a-helices and a cytosolic nucleotide-binding domain that is involved in ATP binding [3]. 113
Figure 5.1. Topology of ABC transporters. The TMD, with the predicted membrane spans and the NBD, containing the nucleotide binding fold, are shown for the three main types of full-size
ABC transporters. Adopted from [4].
Some of the ABC transporters have been characterized in plants; however, a majority of the transporters still remain uncharacterized. A. thaliana consists of about 131 members of the
ABC gene family including 54 full size transporters consisting of two hydrophobic and two hydrophilic domains [5, 6]. Sequencing of plant genomes has allowed us to study the putative
ABC genes and to analyze the evolution of ABC gene family since the divergence of monocots and dicots.
PDR subfamily was identified after observing the drug resistance in S. cerevisiae. PDR5 from yeast was isolated as a transporter mediating resistance to cycloheximide [7], but now
PDR5 has shown resistance to a wide range of structurally and functionally unrelated compounds 114
[8]. Regardless of the structural differences, PDR5 from yeast is considered as a homolog of
MDR1 from human.
PDR transporters are specific to fungi and plants [9]. In plants, the first PDR genes to be identified were SpTUR2 (homolog of yeast PDR5) in the aquaphyte Spirodela polyrrbiza [10] and NpPDR1 in Nicotiana plumbaginifolia [11]. The SpTUR2 transcript is up-regulated by abscisic acid and by environmental stresses, including low temperature, high salt, or cycloheximide [3]. The expression of NpPDR1 has shown to be induced by various microbial elicitors [12], which suggests that some PDR genes might be involved in plant defense.
In Arabidopsis, 15 PDR genes are known out of which 6 PDR genes are extensively studied. AtPDR11 is described as a paraquat transporter localized to the plasma membrane. Due to the structural similarity between paraquat and polyamines, paraquat is recognized as a substrate and is internalized and transported via the polyamine transport system [13]. The interaction between the transportation of polyamines and paraquat has been long reported in plants. In maize roots, paraquat uptake was competitively inhibited by diamines such as putrescine and cadaverine, but was not much affected by spermine and spermidine [14, 15]. In A. thaliana roots; spermidine, spermine and putrescine inhibit the uptake of paraquat [16].
Figure 5.2. Structure of Paraquat
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5.2. Materials and Methods
5.2.1. Phylogenetic Analysis
Protein sequences of all the genes from the PDR family were obtained from TAIR [17] .
The amino acid sequence alignment was created using MUSCLE [18]. Phylogenetic trees were constructed by MEGA 7 [19] using the maximum likelihood method based on the Jones-Taylor-
Thornton (JTT) matrix-based model. The reliability of the trees was tested using a bootstrapping test with 1000 duplicates.
5.2.2. Plant material and growth conditions
A. thaliana ecotype Columbia was used as the wild-type. Homozygous mutants of pdr11
(SALK_067836C) were obtained from the Arabidopsis Biological Resource Center, Columbus,
OH. Plants were grown at 22°C under LD conditions (16 h light/8 h dark).
5.2.3. Phenotypic analysis
Seeds were surface sterilized, vernalized at 4°C for 3 days, geminated on half-strength
MS medium and planted in soil. Plants were grown in a growth chamber at 22°C and a relative humidity of 55%. Stem thickness was measured by using a vernier caliper (Fowler tools and instruments, Boston, USA). Flowering time was measured as the day when the floral stem was one cm above the rosette.
5.2.4. Strains, media and reagents
S. cerevisiae wild type; BY4741 (MATa his3D leu2D met15D ura3D); agp2Δ, a yeast strain deficient in spermidine transport [20]; 22Δ8AA, lacking eight AATs and unable to 116 efficiently use Arg, Asp, Glu, Citrulline, GABA, or Pro [21] were used to characterize the candidate polyamine transporters. The cells were grown in yeast extract-peptone-dextrose (YPD) or synthetic complete (SC) medium at 30C. For PA transport studies, cells were cultured in SC medium with 2 % galactose. [H3] spermidine was obtained from Amersham Biosciences
(Piscataway, NJ, USA). PA and amino acids were obtained from Sigma-Aldrich (St Louis, MO,
USA).
5.2.5. DNA cloning
Full-length cDNA of PDR11 gene was amplified using primers 5’- caccatggccgcaatgctaggacga - 3’ and 5’- ttctcaacttccaaagaaggtga - 3’, it was then cloned into pENTR/D-TOPO cloning vector (Invitrogen, Carlsbad, CA, USA) according to the supplier’s protocol. Subsequently, the target gene was transferred to destination/expression vector pYES-
DEST52 or pAG303 EPD-CCDB by LR recombination reaction (Invitrogen) to generate.
Plasmid DNA was prepared and PCR checked to ensure that the inserts were present.
5.2.6. Functional complementation assay
Wild type, agp2Δ - empty vector and agp2Δ - PDR11 gene transformants were grown in yeast extract-peptone-galactose (YPG) media. The density of exponentially growing cell cultures was normalized to an OD600 of 0.5. Cell suspensions were serially diluted and 3 µl aliquots were spotted onto YPG plates containing 25 mM spermidine. Plates were photographed after 3–4 days of incubation at 30°C. All experiments were repeated two to three times each with three replications.
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5.2.7. Amino acid transport assay
Wild type, 22Δ8AA - empty vector and 22Δ8AA - PDR11 gene transformants were grown in yeast extract-peptone-galactose (YPG) media. Transformants were selected on media lacking uracil supplemented with 10 mM ammonia sulfate. Complementation was performed on
BA minimal media [22] with 1 mM of Arg, Asp, citrulline, GABA, Glu, and Pro as sole nitrogen source. Plates were photographed after 3–4 days of incubation at 30C. All experiments were repeated two to three times each with three replications.
5.2.8. Time course dependent study
Wild type, agp2Δ - empty vector and agp2Δ - PDR11 gene transformants were grown in yeast extract-peptone-galactose (YPG) media. Mid-logarithmic cells were harvested, washed three times with the uptake buffer (50 mM sodium citrate, pH 5.5, 2 % galactose). The cells were then suspended in the uptake buffer and the uptake of polyamines was initiated by the addition of
15µM [H3] spermidine. After 0, 2, 4, 6, 8 minutes respectively, the uptake of polyamines was stopped by adding the uptake buffer containing 150µM [H3] spermidine. The cells were filtered through 0.45 µm Millipore cellulose filter; the filters were then washed with three times with ice cold uptake buffer in order to remove any exogenously attached [H3] spermidine. The filter papers were transferred to scintillation vials containing 10 ml of Ecoscint (National Diagnostics,
Atlanta, GA, USA). The radioactivity on the filters was counted in a Beckman Coulter LS-7000 liquid scintillation counter (Fullerton, CA, USA) after quenching.
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5.3. Results
5.3.1. Phylogenetic Analysis of AtPDR11
Arabidopsis has 15 PDR genes, which can be clustered into 2 major groups. Individual
clusters suggest that during the course of evolution, loss of introns have taken place. PDR11
clusters together with PDR6 and both the genes are expressed in the root. PDR6 and PDR11
differ by an extra intron in the third exon.
Figure 5.3. Phylogenetic analysis of AtPDR protein sequences. Full-length protein sequences were aligned using the Muscle program. Maximum Likelihood method was used to construct the phylogenetic tree with 1000 bootstrap replicates using Mega 7.
119
5.3.2. The tissue/organ expression of AtPDR11 by microarray analysis
Proteins are the major functional components of cells and information about their expression in cells helps us better understand the biology of multicellular organisms. The expression pattern of AtPDR11 was analyzed using the existing microarray database by The Bio-
Analytic Resource for Plant Biology (BAR). The microarray results showed that AtPDR11 had highest expressions in the pedicels, carpels and root.
Figure 5.4. Predicted Tissue/Organ Expression of AtPDR11 (generated with the Plant eFP at bar.utoronto.ca/eplant)
5.3.3. Phenotypic characteristics of AtPDR11 mutant
To analyze whether the mutants of AtPDR11 had different germination rates and root lengths, we germinated the seeds on MS plates and observed the germination rate and the difference in the root length. The germination rate for the wildtype plants were 97.78% whereas for mutants were 62.5%. The mutants of AtPDR11 also had shorter roots as compared to the wildtype plants. 120
It has been reported that changes in the levels of polyamines results in phenotypic alterations in plants. Changes in phenotypic characteristics between wildtype and homozygous mutants of PDR11 were studied. Plants were germinated on MS plates and moved to pots with soil under long day conditions. At four weeks, the mutant plants had already flowered and the wild type plants were delayed for 3-4 days. The mutants also had smaller rosette leaves. After bolting, the flowering stems of the mutants are thinner, than those of the wild-type.
Figure 5.5. Phenotypic Characteristics of wildtype and AtPDR11Δ plants. A, Germination rate of wildtype and homozygous PDR11Δ plants (n = 88). B, Root length of seedlings in wildtype and homozygous PDR11Δ plants. Seedlings were grown for 7 d on one-half strength
MS. C, Comparison of 2.5-week-old wildtype and homozygous AtPDR11Δ plants. D,
Comparison of stem thickness between wildtype and homozygous AtPDR11Δ plants. The diameters of stems were measured in 6-week-old plants 121
5.3.4. Involvement of the AtPDR11 in abiotic stresses
To analyze the physiological role of PDR11 in abiotic stresses, the tolerance level of
PDR11 mutants was assayed. For salt stress and osmotic stress, AtPDR11 mutants were germinated on MS plates and the seedlings were transferred to plates containing 150 mM NaCl,
300mM sorbitol after 7 days. Such an unexpected increase in salinity after germination is common in arid areas that require shift to irrigation with poor water quality. The mutants exhibited a salt sensitive phenotype. The first symptom observed of salt stress in the mutants was bleaching of cotyledons spreading to the true leaves and then resulting in death of the whole seedling. For cold stress, the plates were kept at 4°C for 21 days after germination. Both cold stress and osmotic stress had no effects on the plants.
Figure 5.6. Analysis of salt tolerance in wildtype and AtPDR11Δ plants. A, Survival of wildtype and homozygous AtPDR11Δ grown on 150 mM NaCl. Seedlings were grown for 7 d on one-half strength MS medium and then transferred to 150 mM NaCl for 5 d. B, Number of affected seedlings of wildtype and AtPDR11Δ grown on 150 mM NaCl. 122
5.3.5. Functional characterization of AtPDR11 in yeast
To investigate the potential role of PDR11 in polyamine transport, cDNA of AtPDR11 was amplified and inserted in the yeast expression vector pYES-DEST52. The resulting plasmid construct along with the empty vector pYES-DEST52, were transformed into the yeast AGP2Δ strain. AGP2Δ has a disrupted gene for high affinity spermidine permease and therefore is resistant to high concentrations of polyamines. AGP2Δ can grow in the presence of exogenous polyamines whereas wildtype yeast is sensitive to it. Functional complementation assay shows that the transformants are sensitive to the exogenous polyamines mimicking the phenotype of a wildtype yeast strain. This assay shows that AtPDR11 can transport polyamines.
Figure 5.7. Functional complementation of AtPDR11 in yeast mutant agp2Δ. WT (BY4741), agp2Δ-empty vector and agp2Δ- PDR11 strains were grown overnight on SC medium supplemented with 2% galactose. The density of exponentially growing cell cultures was normalized to an OD600 of 0.5. Cell suspensions were serially diluted as indicated and 3 μl of each were spotted onto YP-galactose plates containing 15 mM spermidine and 25 mM spermidine. Plates were photographed after 3 -4 d of incubation at 30°C. The data is representative of three independent experiments 123
5.3.6. Heterologous expression of the AtPDR11 in yeast
To further understand the transport capabilities of PDR11, cDNA of AtPDR11 was amplified and inserted in the yeast expression vector pAG303 EPD-CCDB, allowing the expression under the control of the GAL1 promoter. The resulting plasmid construct along with the empty vector of pAG303 EPD-CCDB, were transformed into the yeast 22Δ8AA strain.
22Δ8AA lacks eight amino acid transporters and is unable to grow on Arg, Asp, Glu, Citrulline,
GABA, or Pro as the sole N source. Both wildtype and the transformant were able to grow on
BA minimal media in the presence of a single amino acid. Amino acid transport assay shows
AtPDR11 can transport amino acid from the BA minimal media and use it as a sole N source to survive.
124
Figure 5.8. Amino acid transport assay of AtPDR11 in yeast mutant 22Δ8AA. Cell suspensions were streaked onto BA minimal media with Arg, Asp, Glu, Citrulline, GABA, or
Pro. Plates were photographed after 3 -4 d of incubation at 30°C.
5.3.7. Transport of polyamines by AtPDR11
To determine the ability of PDR11 to transport polyamines, radiolabeled spermidine was used to conduct a transport experiment. 15µM of [H3] spermidine was incubated with wildtype, empty vector and the transformants for 0, 2, 4, 6 and 8 minutes respectively to determine the uptake of radiolabeled spermidine. In wildtype, the uptake of [H3] spermidine was highest over the first 4 minutes and then the uptake started declining. The PDR11 transformant showed increased uptake of [H3] spermidine compared to the empty vector. At 8 minutes, there was no difference between the uptake by the wildtype and the transformant.
12000
10000
8000
DEST52-AGP2Δ 6000 WT PDR11 4000
2000
0 2 4 6 8
Figure 5.9. Time course uptake of [H3] spermidine. Log phase cells were incubated with
[14C] spermidine for 0, 2, 4, 6 and 8 min. 125
5.4. Discussion
This study shows that AtPDR11 is a broad specific transporter and can transport polyamines as well as amino acids. Previous studies have shown that the uptake of paraquat is competitively inhibited by polyamines, indicating that both polyamines and paraquat are transported via the same transport system [14, 23]. PDR11 was described as a paraquat transporter suggesting the possibility of it transporting polyamines as well.
Yeast assay shows that AtPDR11 can take up exogenous polyamines and mimic the wildtype phenotype, suggesting that PDR11 can transport polyamines (Figure. 7). We further studied the transport capability of AtPDR11 to transport polyamines. The time course uptake of
[H3] spermidine shows that AtPDR11 can take up labeled polyamines as well (Figure. 9).
Most of the rice ABC genes identified have at least one A. thaliana homolog to which they share more than 60% amino acid identity, this reveals that gene duplication or deletion must have occurred after the divergence of monocots and dicots [24]. The phylogenetic tree of PDR11 proteins from A. thaliana shows AtPDR11 is closely related to AtPDR6 (Figure.3). AtPDR11 is sensitive to salt stress (Figure.6) and there are certain similarities which exists between the functionalities of both genes regarding the response to biotic stress [25, 26]. The polyamine transporter mutant pdr11 flowered 4 days earlier and had thinner stems when compared to the wildtype. The mutants also exhibited lower germination rates and smaller roots (Figure. 5).
The amino acid transporter is expressed in roots (Figure.4). Recently, four amino acid transporters, LHT1, AAP1, AAP5 and ProT2 have been shown to play vital role in the uptake of amino acid via root [27, 28, 29, 30, 31]. Amino acid transport assay shows that AtPDR11 can transport amino acids and use a single amino acid as the sole source for nitrogen (Figure. 8). 126
Although for information of whether the transporters are directly or indirectly involved in the import of amino acids through roots, uptake studies need to be performed using plants.
In A. thaliana, the translocation of amino acid from source to sink takes place in the phloem and requires the loading of organic nitrogen in the minor veins of the leaf [32]. However, the transporters required for this have not yet been identified, but potential candidates might belong to the AAP family [33].
127
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791, 2012.
CHAPTER VI. 134
FUNCTIONAL CHARACTERIZATION OF A VACUOLAR POLYAMINE
TRANSPORTER
6.1. Introduction
OCTs (Organic cation transporters) are termed as uniporters and belong to major facilitator superfamily (MFS). Based on their amino acid sequence OCTs have been assigned to
SLC22A family. OCTs are responsible for mediating the transport of organic cations in and out of the cell through facilitated diffusion [1].
In plants, the first OCT to be studied was Phaseolus vulgaris OCT1 (PvOCT1), a 547 amino acid protein predicted to have twelve membrane spanning domains which are higly conserved in the MFS [2]. Northern analysis showed that PvOCT1 is strongly expressed in roots and stems, while in situ hybridization revealed the existence of PvOCT1 in phloem cells.
Dehydration stress exponentially increased the transcript levels of PvOCT1 in roots followed by a sudden decrease.
A homology search of this protein with other plants revealed that it had a strong similarity ranging from 51% to 71% with six proteins of A. thaliana. In A. thaliana, the first protein to be characterized as OCT was AtOCT1 which was localized to the plasma membrane
[3]. The other five OCTs - OCT2-6 were localized to the tonoplast [4].
Like PUTs, OCTS are also involved in playing pivotal roles against environmental stresses. During drought stress, AtOCT4, AtOCT5 and AtOCT6 are up-regulated; during cold stress, AtOCT3 and AtOCT5 are up-regulated and during salt stress treatments AtOCT 5 and 6 are up-regulated. On the other hand, AtOCT1 does not respond to these stresses [4]. 135
In humans and rodents, OCT 1 and OCT 2 are expressed in kidney and liver and are involved in the renal elimination of xenobiotic compounds. Transport of [3H]agmatine and
[3H]putrescine in human embryonic kidney (HEK293) cells stably transfected with hOCT1, hOCT2 showed concentration-dependent agmatine transport. hOCT2-HEK cells also showed pH- and concentration-dependent putrescine accumulation.. [5, 6]. In rats, electrogenic transport shows OCT1 can transport spermine and spermidine [7]. Spermidine, which is longer and carries an additional positive charge, is also a transported substrate of mammalian OCT1, OCT2, and
OCT3 [8].
Predicted membrane topology of OCT1 from rats consists of 12 a-helical transmembrane domains (TMDs), a large extracellular loop between TMDs 1 and 2 and a large intracellular loop between TMDs 6 and 7. Predicted membrane topology of OCT5 from A. thaliana, have a similar structure as that of OCT1 from rats and this consists of 11 helical TMDs with a large extracellular loop between TMDs 1 and 2 and a large extracellular loop between TMDs 6 and 7.
Figure 6.1. Predicted membrane topology of OCT1 from rats and OCT5 from A.thaliana. 136
OCTs are also known to transport Metformin, the most widely prescribed drug for the treatment of Type II diabetes [9]. In rats, Metformin is transported by OCT1. In Mice, OCT1 is responsible for the hepatic and intestinal uptake of metformin and in humans; both OCT1 and
OCT2 are responsible for the transport of metformin. OCTs are also involved in the transportation of various different exogenous and endogenous substances such as TEA, MPP cimetidine, monoamine neurotransmitters, acetylcholine, histamine [10] .
Table 6.1. Structure comparison of putrescine, spermidine, spermine and metformin.
Based on the research carried out in other organisms, and taking into consideration that metformin is a structure analog of polyamine we hypothesized that OCTs play a role in the transportation of polyamines in plants. We decided to characterize OCT5 based on its involvement in different stress responses. Organ-specific expression reveals the expression of
OCT5 in sink leaves and source leaves and tissue-specific expression shows the expression of it in leaves, siliques, in roots, stems, and flowers [4].
6.2. Materials and Methods
6.2.1. Plant material and growth conditions 137
A. thaliana ecotype Columbia was used as the wild-type. Homozygous mutants of OCT5
(SALK_045610C) were obtained from the Arabidopsis Biological Resource Center, Columbus,
OH. Plants were grown at 22°C under LD conditions (16 h light/8 h dark).
6.2.2. Phenotypic analysis
Seeds were surface sterilized, vernalized at 4°C for 3 days, germinated on half-strength
MS medium and planted in soil. Plants were grown in a growth chamber at 22°C and a relative humidity of 55%. Stem thickness was measured by using a vernier caliper (Fowler tools and instruments, Boston, USA).
6.2.3. Polyamine toxicity assayBY4741 (wild type yeast), AtOCT5- BY4741 (overexpressed yeast construct), TPO5 Δ (mutant of a spermidine and putrescine exporter that is localized to the
Golgi [11]) and AtOCT5-TPO5 Δ (complementation) were grown in CSM-GAL until OD600 =
0.2, after which the cell growth was measured in the presence of 3, 6 and 9 µM of spermidine by using a microplate reader.
6.2.4. Subcellular Localization of OCT5
Full-length cDNA sequence of AtOCT5 was cloned into the GFP expression vector pGWB5 using the GATEWAY® recombination system [12]. Inserts were verified by using
PCR, and vectors were transformed into Agrobacterium tumefaciens strain GV3101 [13]. The mCherry-Vacuolar Marker (CD3-976) [14] was obtained from ABRC and used as an organellar marker. Images of one-micron sections through the leaf were acquired using a Leica TCS SP5 multi-photon laser scanning confocal microscope at 12-72 hours after infiltration.
6.3. Results 138
6.3.1. Phenotypic characteristics of AtOCT5 mutant
To analyze whether the mutants of AtOCT5 had different germination rates and root lengths, we germinated the seeds on MS plates and observed the germination rate and the difference in the root length. No differences were observed in the germination rates and the root lengths between the wild type plants and the mutant plants.
We also studied the differences in the phenotypic characteristics between wild-type and homozygous mutants of OCT5. Plants were germinated on MS plates and moved to pots with soil under long day conditions. The mutants of OCT5 plants were a bit smaller when compared to the wild-type plants.
Figure 6.2. Phenotypic Characteristics of wildtype and AtOCT5Δ plants. A, Comparison of
5-week old wild-type and homozygous AtOCT5Δ plants. B, Root length of seedlings in wild- type and homozygous OCT5Δ plants. Seedlings were grown for 7 d on one-half strength MS. C, 139
Comparison of stem thickness between wild-type and homozygous AtPDR11Δ plants. The diameters of stems were measured in 6-week-old plants.
6.3.2. Involvement of the OCT5 in abiotic stresses
We further analyzed the role of OCT5 mutant plants in abiotic stresses, the tolerance level of OCT5 mutants were assayed. For salt stress and osmotic stress, AtPDR11 mutants were germinated on MS plates and the seedlings were transferred to plates containing 150 mM NaCl,
300mM sorbitol after 7 days. Such an unexpected increase in salinity after germination is common in arid areas that require a shift to irrigation with poor water quality. The mutants exhibited a salt-sensitive phenotype (Figure. 3). For cold stress, the plates were kept at 4C for 21 days after germination. Both cold stress and osmotic stress had no effects on the plants.
Figure 6.3. Analysis of salt tolerance in wild-type and AtOCT5Δ plants. A, Survival of wild- type and homozygous AtOCT5Δ grown on 150 mM NaCl. Seedlings were grown for 7 d on one- 140 half strength MS medium and then transferred to 150 mM NaCl for 5 d. B, Number of affected seedlings of wild type and AtPDR11Δ grown on 150 mM NaCl.
6.3.3. Polyamine toxicity assay in S. cerevisiae:
To determine the effect of exogenous polyamines on yeast, BY4741, AtOCT5- BY4741,
TPO5 Δ (mutant of a protein involved in excretion of putrescine and spermidine) and AtOCT5-
TPO5 Δ were grown in varying amounts of spermidine. All the yeast constructs started off at
OD600 = 0.2 (Figure 4.A) but the presence of exogenous spermidine inhibited the growth of
TPO5 Δ resulting in compromised growth which is followed by BY4741, AtOCT5- TPO5 Δ and
AtOCT5- BY4741 (Figure. 4). At 9 µM, AtOCT5- BY4741 is resistant to polyamine toxicity
(Figure. 4.D). This shows that OCT5 is involved in the excretion of polyamines from the cells.
141
Figure 6.4. Polyamine toxicity assay in S. cerevisiae. A, TPO5. B, Wildtype. C, OCT5-TPO5.
D, Oct5-Wildtype.
6.3.4. Subcellular localization of OCT5
Confocal analysis of GFP-OCT5 fusion construct shows a compartmental localization within the Vacuole. Co-expression with an mCherry Vacuolar-marker confirms the GFP localization to the Vacuole in the leaves of N. benthamiana (Figure. 4).
Figure 6.5. Sub-cellular localization of AtOCT5. A, GFP signal. B, mCherry – Vacuolar
Marker. C, Chloroplast autofluorescence. D, Merged image. 142
6.4. Discussion
This study shows that AtOCT5 is a vacuolar protein, sensitive to exogenous polyamines.
Like Polyamine Uptake Transporters (PUTs), that are highly expressed in various stress responses in plants OCTs are also known to have been involved in different stress responses such as drought stress, cold stress, and salt stress. OCT5 is highly expressed in these stress conditions; therefore, we chose to characterize OCT5.
Previous studies have shown that in mammals OCTs are involved in the transportation of putrescine, agmatine, spermine and spermidine [15, 5, 16]. In mammals, OCTs are also known to transport metformin which is structurally similar to the polyamines and contains multiple amino groups (Table 1).
Since polyamines are essential for the growth and development of plants and play important roles in various plant processes such as root formation, senescence, fruit ripening, etc.
[17, 18, 19], and therefore transportation of polyamines also affects the growth of a plant. We studied the phenotypic characteristics of a mutant of OCT5 and wildtype, the mutant plants were smaller than that of a wild-type showing the lack of transportation directly affects the growth and hinders growth (Figure. 2). We also performed abiotic stress experiments and observed that mutants of OCT5 are sensitive to salt stress (Figure.3), however, we did not observe differences in cold stress and osmotic stress. The experimental protocol was different from that of the Küfner
& Koch [4].
Although polyamines are essential for both prokaryotes and eukaryotes, excessive polyamines can be toxic for the organism if it is not used or excreted. Confocal microscopy of
Nicotiana leaves filtrated with GFP fused OCT5 constructs reveals the sub-cellular expression of it in vacuoles (Figure. 5). Vacuoles are organelles responsible for the storage and/or excretion of 143 wastes from the cell. Therefore the presence of exogenous polyamines is toxic for a mutant of vacuolar polyamine transporter, TPO5 (Figure. 4). Whereas overexpression construct OCT5-WT grow without any negative effect in the presence of exogenous polyamines.
Different polyamine transporters have been shown to be localized to different parts of the plants inferring that polyamines are present globally in the cell. PUT2 and PUT3 are localized to the chloroplast, PUT5 is localized to the ER and PDR11 are localized to the plasma membrane.
To better understand the function of OCT5 as a polyamine transporter, further transport experiments are needed. HPLC analysis of the yeast construct used in the toxicity experiment referring to a mutant, wildtype, functional complementation and overexpression; and of mutant plants of OCT5 and wildtype will shed more light over the polyamine contents. These experiments will demonstrate the transport capabilities of OCT5 to transport polyamines.
144
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[10] J. Johan and S. Alfred, "Pharmacological and Physiological Functions of the Polyspecific
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148
CHAPTER VII.
SUMMARY
Polyamines are essential metabolites that help regulate almost all the major cellular processes of plants. However, the molecular mechanisms behind these are not known yet. The intracellular levels of polyamines are regulated by biosynthesis, catabolism, conjugation and transport. Although extensive studies have been done on the metabolism of polyamines, most of the protein involved in polyamine transport and metabolism have not been characterized. This thesis has made four significant contributions to plant polyamine biochemistry.
7.1. Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways and the effect of high temperature on the delay of flowering in PUT5 plants
Polyamines have effects similar to hormones in that they are transported throughout the plant, and changes in their levels are associated with processes such as embryogenesis, fruit development, ripening, senescence, tissue responses to biotic and abiotic stresses. An essential function of spermidine is that it is the substrate needed for the hypusination of a conserved transcription factor elf5A in all eukaryotes. Inducible suppression of deoxyhypusine synthase results in early flowering, and RNAi suppression of deoxhypusine synthase using a leaf–specific promoter does not affect flower time, but promotes leaf growth and increases the number of flowers. Here we show that expression of spermidine-preferential transporters PUT5:OsPUT1 and CaMv:OsPUT1 in A. thaliana results in plants with larger leaves, thicker stems, and a delay of flowering of 14-16 days under permissive flowering conditions. In contrast, the homozygous
KO mutants of AtPUT5 flower 3-4 days earlier than WT plants, and also have smaller leaves and 149 wispier stems. The delayed flowering phenotype is correlated with elevated levels of spermidine and spermidine conjugates in the leaves of the transgenic plants. CaMv:OsPUT1 but not
PUT5:OsPUT1 plants were also significantly delayed in senescence. Transient expression of both GFP-tagged AtPUT5 and OsPUT1 in N. benthamiana shows that these transporters are localized to the ER. The delay of flowering in the transgenic plants disappear when gibberellins are applied externally or when the plants are grown at ~28 . This shows that the delay of flowering phenotype due to the overexpression of a polyamine℃ transporter is present upstream of the regulation of flowering due to the gibberellin response pathway or the temperature response pathway.
7.2. Dual functioning of plant arginases provides a third route for putrescine synthesis
In plants, putrescine is synthesized via ornithine decarboxylase pathway (ODC) and the arginine decarboxylase pathway (ADC). However, over the course of evolution, A. thaliana has lost its ODC and has two ADC genes. ADC1 participates in the ADC pathway for the synthesis of putrescine and is localized to the cytoplasm. Here we show that there exist a second putrescine biosynthetic pathway localized to the chloroplast which can synthesize putrescine with the help of ADC2 and ARGAH2. ARGAH have dual specificity and can function as arginase or agmatinase depending on the concentration of arginine. Yeast complementation assay reveals that ADC2 and ARGAH2 can successfully synthesize putrescine via arginine. HPLC analysis of
ADC2 and ARGAH2 conducted in the presence of 1mM levels of arginine show accumulation of agmatine and putrescine. This results reveal that at 1mM levels of arginine, ARGAH functions as agmatinase. Sub-cellular localization and functional complementation assay shows the existence of the chloroplast localized pathway in Glycine max as well. To further understand the function of this pathway in other plants, we studied the phytozome database which shows the possibility 150 of the existence of this pathway in four members of the Brassicaciae family, S. lycopersicum and
G. max. Since we noted spatial separation of the two putrescine biosynthesis pathways starting with ADC, we postulated that synthesis of ODC might be spatially separated as well. The sub- cellular localization of ODC from O. sativa and Glycine max reveals that plants which have retained their ODC have their ODC localized to the ER. This study will certainly help us in understanding the distribution of polyamines in the different compartments of a plant cell.
7.3. PDR11 is a broad substrate transporter of polyamines and selected amino acids
In Arabidopsis, the ABC transporter PDR11 was first described as a plasma membrane- localized paraquat transporter. Since paraquat is known to be transported by polyamine transporters, we hypothesized that PDR11 was also a polyamine transporter. To test this hypothesis, the full-length cDNA was expressed in the yeast mutant AGP2∆ which is deficient in polyamine transport. Yeast functional assays demonstrated time-dependent uptake of radiolabeled [H3] spermidine. Significant uptake rates were noted at 15 µM levels of spermidine, indicating that this transporter has a high affinity for spermidine transport.
Homozygous mutants of PDR11 resulted in lower germination rates, smaller roots, smaller rosette leaves, thinner stems and a delay in the timing of flowering when compared to the wild-type plants. Similar to most polyamine transporters, PDR11 also exhibit sensitivity to abiotic stress. To test the hypothesis that PDR11 might also transport other nitrogenous compounds, PDR11 was transformed and expressed in in the yeast amino acid transport mutant strain mutant strain 22∆8AA. Complementation assays show that heterologous expression of
PRD11 will support the growth of this mutant strain when any of arginine, aspartic acid, 151 glutamic acid, citrulline, GABA, or proline is added to the medium. Thus, PDR11 is a polyamine transporter with the ability to transporter at least some amino acids.
7.4. Functional characterization of a vacuolar polyamine transporter
In mammals, OCTs are involved in the transportation of putrescine, agmatine, spermine, spermidine and a structurally similar molecule known as metformin. Therefore, we hypothesized that OCT5 can also transport polyamine. We studied the phenotypic characteristics of a mutant of OCT5 and wildtype, the mutant plants were smaller than that of a wild-type. Like most of the polyamine transporters, OCT5 are sensitive to salt stress. Confocal microscopy of N. benthamiana leaves filtrated with GFP fused OCT5 constructs reveals the sub-cellular expression of it in vacuoles. Vacuoles are organelles responsible for the storage and/or excretion of wastes from the cell. Yeast polyamine toxicity assay reveals that the presence of exogenous polyamines is toxic for a mutant of vacuolar polyamine transporter, TPO5. Whereas overexpression construct grow without any negative effect in the presence of exogenous polyamines. This study shows that AtOCT5 is a vacuolar protein, sensitive to exogenous polyamines.
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https://s100.copyright.com/CustomerAdmin/PLF.jsp?ref=29decb3c-f473-41b0-85ab-67ce3bd9aaa2 6/6 Dear Alexander: This letter indicates that I am giving permission to have Sheaza Ahmed include in her dissertation, two papers where I am listed as the senior author.
Best Regards Paul
Paul Morris Professor, Biological Sciences Program Director, Kids' Tech University http://personal.bgsu.edu/~pmorris/Site/Welcome.html http://kidstechuniversity-bgsu.vbi.vt.edu/program.php Bowling Green State University Bowling Green, OH [email protected] 419 372 0481 Bowling Green State University Bowling Green Ohio
Sheaza Ahmed Bowling Green State University Department of Life Sciences 217 Life Sciences Building Bowling Green, OH 43403
September 18, 2017
RE: Permission for publication of our journal articles in your dissertation
Dear Sheaza,
I give you permission to publish the jour al articles Altered e pressio of pol a i e tra sporters re eals a role for sper idi e i the ti i g of flo eri g a d other de elop e tal respo se path a s a d Dual fu ctio i g of pla t argi ases pro ides a third route for putresci e s thesis as a part of our dissertation.
Sincerely,
Andrea Kalinoski, Ph.D.
Andrea L. Kalinoski, Ph.D.
Associate Professor · Technical Director, Integrated Core Laboratories · Department of Surgery · University of Toledo College of Medicine and Life Sciences · 3000 Arlington Avenue· Toledo, OH 43614 · Phone 419-383-4205 · Fax 419-383-6230 [email protected]
Hi Alex, I give the permission to publish our journal articles “Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways” and “Dual functioning of plant arginases provides a third route for putrescine synthesis” as a part of the dissertation of Sheaza Ahmed, Department of Life Sciences, Bowling Green State University.
Thank You, Vipa Phuntumart —
Vipaporn Phuntumart, Ph.D. Associate Professor 129 LifeSciences Building Department of Biological Sciences Bowling Green State University Bowling Green, OH 43403 Phone: 419-372 4097 Fax: 419-372 2024 http://www.bgsu.edu/arts-and-sciences/biological-sciences/faculty-and-staff/alphabetical-listing/vipa-phuntumart.html
September 20, 2017 Sheaza Ahmed Department of Life Sciences Bowling Green State University
RE: Permission for publication of our journal articles in your dissertation
Dear Sheaza,
I give you the permission to publish our journal articles “Dual functioning of plant arginases provides a third route for putrescine synthesis” as a part of your dissertation.
Sincerely, Lingxiao Ge September 19, 2017 Sheaza Ahmed Department of Life Sciences Bowling Green State University
RE: Permission for publication of our journal articles in your dissertation
Dear Sheaza,
I give you the permission to publish our journal articles “Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways” and “Dual functioning of plant arginases provides a third route for putrescine synthesis” as a part of your dissertation.
Thank You, Menaka Ariyaratne September 18, 2017 Sheaza Ahmed Department of Life Sciences Bowling Green State University
RE: Permission for publication of our journal articles in your dissertation
Dear Sheaza,
I give you the permission to publish our journal articles “Altered expression of polyamine transporters reveals a role for spermidine in the timing of flowering and other developmental response pathways” and “Dual functioning of plant arginases provides a third route for putrescine synthesis” as a part of your dissertation.
Thank You, Jigar Patel