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INTRODUCTION

1.1 Stressors

Substantial loss of yield worldwide is due to abiotic stress (Shelden and

Roesnner 2013). In fact, more than 50% of the yield is lost to abiotic stress

(Boyer, 1982, Bray et al. 2000 as stated in Wang et al. 2003, Shelden and Roesnner

2013). For instance, in wheat, the record yield (average yield + yield without

loss to diseases, pests, and unfavouable environments) in 1975 was 14500

kg/hectares while the average yield was 1880 kg/hectares; further, losses to

unfavourable physicochemical environments was 12156 kg/hectares (Boyer, 1982).

Translated, this would mean a loss of 87% to diseases, pests, and unfavourable

physicochemical environments; in particular, a loss of 71.7% to unfavourable

environmental conditions (Boyer, 1982).

Drought, salinity, cold, heat, oxidative stress, and chemical toxicity are the

primary stressors of a plant (Carson 2012, Wang et al 2003). The damage and

cellular response of plant to any of the above stressors tend to be similar;

they are interconnected in terms of the response induced by the plant (Wang et

al. 2003). However, cell identity and the number of simultaneous stresses

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determine stress-induced gene expression patterns (Dinneny et al. 2008 as

stated in Voesenek and Pierik, 2008). When both heat and drought stresses were

simultaneously subjected onto a plant – Arabidopsis thaliana – only 10% of the

genes imbricated between the double-stressed and a single-stressed treatment

were expressed (Voesenek and Pierik, 2008). The latter would imply that the

gene expression profile, and control networks of multiple stresses cannot be

adumbrated by expression profiles generated for single stress treatments

(Voesenek and Pierik, 2008).

Stressors usually cause disruption of osmotic and ionic homeostasis, or damage

to functional, and structural proteins, and membranes (Wang et al. 2003, Figure

1). The genetic makeup of any given plant is responsible for the stress-induced

response. However, acclimation can also influence the response of a plant

(Carson 2012). Genes regulated by the stressors usually are involved in

signaling cascades, or transcriptional control, function to directly protect

membranes and proteins, or are involved in water and ion uptake and

transportation (Wang et al. 2003). Tolerance to stressors is usually conferred

by modifying the internal environment to permit internal structural

stability, and function of internal components (Wang et al. 2003).

In other words, if one can manage to retain the structural integrity of the

cell, the effect of the stressor would be minimized (Wang et al. 2003). Response

to salt stress usually involves tolerance (plant maintains water uptake, and

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growth), Na+ exclusion (removal of toxic ions from shoot tissue), and tissue

tolerance (selectively isolate toxic ions into vacuoles (Munns and Tester, 2008

as stated in Shelden and Roesnner 2013). Munns and Tester (2000, 2008) (as stated

in Shelden and Roesnner 2013) noted that a sub-population of the duram wheat

genotypes possessed the ability to tolerate salt stress via Na+ exclusion;

later, Nax1 and Nax2 were identified as the genes conferring this tolerance,

and incorporated into alternate lines to decree salt-stress tolerance to scions

(Shelden and Roesnner 2013).

In Triticum aestivum, for example, a number of genes were found to be up-

regulated by the said stressors (Khalil 2013, Tardif et al. 2007). A scad of these

genes are regulatory factors (ex: protein kinases, transcription factors,

ubiquitin ligases, GTP and calcium binding proteins; therewith, up-regulated

genes may act to modify chromatin or act as post-transcriptional regulation

(Tardif et al. 2007). However, the external architecture of the plant is also

retailored to specific stresses. For instance, in response to water stress,

plants have been noted to suppress lateral root formation, and initiate

primary root formation in an attempt to reach the deeper layers of the soil

where the probability of water discovery is superlative (Xiong et al 2006 as

stated in Wright 2014, thesis).

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Figure 1: Mechanisms of stress tolerance and resistance. (Source: Wang et

al. 2003)

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1.2 Heterotrimeric G Proteins

Animals

G proteins can be classified into two distinct categories – the heterotrimeric

G proteins, and the monomeric G proteins (the small G proteins); the monomeric G

proteins do not include the Gsβγ subunits. However, both the receptor-binding

region and GTPase activity are found on the Gsα subunit; in other words, absence

of either function on the Gsβγ subunit permits exclusion of said subunit by the

monomeric G protein complex.

The heterotrimeric G proteins are involved in signal transduction pathways.

Coupled with cell-surface receptors (henceforth referred to as the G-protein

coupled receptors (GPCR)), the Gs proteins are a network of three discrete

subunits – the α, β, and γ subunits (Lodish et al., 2000, Figure 2). Ligand

binding induces the dislocation and dissociation of the Gsα subunit. The entire

complex vacillates between two alternate states – the “on”, and the “off” state.

This vascillation is dependent upon the bound state of the Gsα subunit. In other

words, in the “off” state, the Gsα subunit is bound to a GDP molecule. However,

ligand binding to GPCR dislodges the GDP and permits a GTP molecule to bind the

α subunit – henceforth referred to as the Gsα GTP complex. The dislocated α

subunit then activates an effector protein which activates an intercellular

second messenger (Lodish et al., 2000). Simultaneously, the bound GTP is

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hydrolyzed, and the Gsα subunit returns to the initial heterotrimeric state,

bound to the β, and γ subunits.

Plants

In plants, only five GPCRs have been found to interact with the Gsα subunit –

AtGCR1, AtRGS1, GPCR, and GTG1, and GTG2 (REF---paper, Khalil et al. 2011).

Effector proteins, on the other hand, can be any one of the following: Ca2+,

calmodulin (CaM), cAMP, cGMP, cADP-ribose, inositol 1,4,5-triphosphate (IP3) and

reactive oxygen species (ROS) (Tardif et al. 2007). The signaling cascade can

result positively or negatively regulate expression of target genes (Tardif et

al. 2007).

Response to stress – pathogens, light, high salinity, hypoxia, drought, ethylene

signaling, signaling pathways regulated by jasmonic acid, gibberellin and

abscisic acid (ABA) - in plants has been noted to involve G-protein signaling.

Plant response to pathogens, for instance, is elicited by modulating

intracellular Ca2+ levels which is itself triggered by G proteins.

Animals possess 23 Gα, 6 Gβ, and 12 Gγ encoding genes while plants possess one

Gα (GPA1), one Gβ (AGB1), and 2 Gγ (AGG1, AGG2) encoding genes (Jones 2002, Figure

2). In plants, the subunits are designated as follows: GPA1 (alpha subunit), AGB1

(beta subunit), AGG1/2 (gamma-A subunit), and AGG3 (gamma-C subunit). G proteins

are involved in regulation of ion-channels, seed germination, light responses,

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cell division, cell elongation, and phytohormones (ABA, GA, auxin) (Jones and

Assmann 2004). In particular, heterotrimeric G proteins in plants have been

noted to induce phytohormones, and control mitosis (Jones 2002). Mutations in

Arabidopsis GPA1 leads to fewer mitosis during formation of hypocotyl, and

leaf; overexpression leads to ectopic mitosis, and overproliferation of

meristem suggesting that GPA1 is involved in the control of cellular division

(Jones 2002).

G proteins in plants are believed to function in ion channel, ABA signaling,

and cellular proliferation (Tuteja and Sopory, 2008). Mutations in members of

G-protein signaling have been shown to affect seed germination, shoot/root

growth, and control of stomata (Tuteja and Sopory, 2008). The G-protein system

in plants is still under study as not much is known about the system (Tuteja

and Sopory, 2008).

Animals have about 1000 GPCRs while plants have been noted to have a single

GPCR (Tuteja and Sopory, 2008). In Arabidopsis thaliana, a GPCR called GCR2 was

found for ABA, however, ABA is recognised by a number of plant receptors

(Tuteja and Sopory, 2008). In plants, there are three known effectors -

Phospholipase D (PLD), K+ and Ca2+ channels (Jones 2002). PLD converts

phospholipids to phosphatidic acid (PA) – where PA acts as a second messenger -

which activates genes involved in stress tolerance, defense, meiosis,

phytochrome action, membrane metabolism, and trafficking of vesicles (Tuteja

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and Sopory, 2008).

Figure 2: Proposed model of stress-induced G-protein signaling cascade in

animals versus plants. (Source: Tuteja and Sopory 2008)

1.3 RD20/Clo3

One stress-induced gene is CLO3/RD20 – which was found to increase during

stresses (Wright 2014). Ta-CLO3 (CLO3 in Triticum aestivum) was noted to

interact with Ta-GA1 – the Galpha subunit in Triticum aestivum (Wright 2014).

In Arabidopsis thaliana, under abiotic stresses, a homolog of CLO3 was found to

be activated; a member of the caleosin family, CLO3 encodes a protein that is

highly abundant when subjected to abiotic stresses. ABA as well as other

stresses (water, salt, osmotic) were found to increase CLO3/RD20 in Arabidopsis

thaliana – a 150 fold increase in CLO3 after 2 hours of dehydration, and a 300

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fold increase 3 hours after ABA treatment (Aubert et al. 2010). In particular,

CLO3 was found to modulate stomatal opening/closing in guard cells (Aubert et

al. 2010). RD20 has been noted to be induced by water stress, ABA, and NaCl

treatments; RD20 was not found to be upregulated by cold treatments (Takahashi

et al., 2000). RD20 mRNA can be observed in leaves, flowers, stems, and siliques,

but not in roots or mature seeds (Takahashi et al., 2000).

1.4 Bimolecular Fluorescence Complementation (BiFC)

Bimolecular fluorescence complementation (BiFC) is a method utilized to detect

protein-protein interactions in vivo. Two potential interacting partners –

the bait, and the prey – are non-fluorescing. Fusion proteins of the bait, and

prey are created in such a manner that the N-terminal of a fluorescent

protein (ex: YFP) is fused to one partner while the C-terminal is fused to the

other (Kerppola, 2008). Interaction between bait, and prey brings into close

proximity the two halves of the fluorescent proteins – a bimolecular

fluorescence complex is formed - which if excited, emits fluorescence

(Kerppola, 2008). BiFC can be used to identify the subcellular location of a

protein complex (Kerppola, 2008).

The advantage of using BiFC is that its intrinsic signal is strong enough to

enable detection; alternative methods require exogenous chromogenic or

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fluorogenic agents (Kerppola, 2008). Further, a number of interactions can be

detected simultaneously by using distinct fluorescence proteins (ex: YFP, CYF,

etc…). However, the signal is usually detected after the formation of the

complex – there is a time lapse between actual interaction, and visibility of

observation, and is therefore not considered a real-time observation (Kerppola,

2008).

Given that not all fluorescent proteins are candidates for BiFC, two in

particular have been noted to function well – YFP, and CFP. YFP is usually

truncated in the following manner: 1-154 consists of the N-terminal portion,

and 155-238 consists of the C-terminal portion; alternatively, one can generate

an N-terminal from 1-172, and a C-terminal from 173-238 (Kerppola, 2008,

Figure 3). Further if the concentration of the fluorescent proteins is high

within a cell, random association between fragments can generate false

positives; it is therefore recommended that fusion proteins be expressed at low

levels or levels typical to the cell (Kerppola, 2008). BiFC usually requires

multiple attempts or generation of multiple combinations (FIG) to determine the

combinations that [work], and that generate the strongest signal (Kerppola,

2008).

BiFC also requires negative controls to validate the protein-protein

interaction detected. Efficiency is calculated as the ratio of the fluorescence

emitted by the YN-YC complex to an internal control (ex: CFP) (Kerppola, 2008).

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Figure 3: Combination of fusion proteins. (Where YN = N-terminal of the YFP

fluorescent protein; YC = C-terminal of the YFP fluorescent protein) (Source:

(Kerppola, 2008))

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References

Aubert, Y., Vile, D., Pervent, M., Aldon, D., Ranty, B., Simonneau, T., Vavasseur,

A., and Galaud, J. (2010). RD20, a Stress-Inducible Caleosin, Participates in

Stomatal Control, Transpiration and Drought Tolerance in Arabidopsis

thaliana. Plant And Cell Physiology 51, 1975-1987.

Boyer, J. (1982). Plant Productivity and Environment. Science 218, 443-448.

Carson, R. (2012). The Interaction of the Stress Regulated Wheat (Triticum

aestivum) Proteins, ESI2 and RAN1, and the Localization of Ipp-1 and Pr-1.

Master of Science. Concordia University.

Nallamsetty, S., and Waugh, D. (2007). A generic protocol for the expression

and purification of recombinant proteins in Escherichia coli using a

combinatorial His6-maltose binding protein fusion tag. Nature Protocols 2,

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Lodish, H., Berk, A., Zipursky, L., Matsudaira, P., Baltimore, D., and Darnell, J.

(2000). Molecular Cell Biology, New York.

Jones, A. (2002). G-protein-coupled signaling in Arabidopsis. Current Opinion

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Jones, A., and Assmann, S. (2004). Plants: the latest model system for G-protein

research. EMBO Rep 5, 572-578.

Kerppola, T. (2008). Bimolecular Fluorescence Complementation (BiFC)

Analysis as a Probe of Protein Interactions in Living Cells. Annual

Review of Biophysics, 37(1), pp.465-487.

Khalil, H. (2013). The characterization of the caleosin gene family in

Triticeae and their role in G-protein signalling and Identification

and characterization of rye genes silenced in allohexaploid triticale:

A bioinformatic study. Doctor of Philosophy. Concordia University.

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Tardif, G., Kane, N., Adam, H., Labrie, L., Major, G., Gulick, P., Sarhan, F., and

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