THE ROLE OF KEA1 AND KEA2 TRANSPORTERS IN PLASTID ION HOMEOSTASIS

AND GENE EXPRESSION

By

RACHAEL ANN DETAR

A dissertation submitted in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

WASHINGTON STATE UNIVERSITY Program in Molecular Plant Sciences

MAY 2021

© Copyright by RACHAEL ANN DETAR, 2021 All Rights Reserved

© Copyright by RACHAEL ANN DETAR, 2021 All Rights Reserved

ii

To the Faculty of Washington State University: The members of the Committee appointed to examine the dissertation of RACHAEL

ANN DETAR find it satisfactory and recommend that it be accepted.

Hans-Henning Kunz, Ph.D., Co-Chair

Helmut Kirchhoff, Ph.D., Co-Chair

Kiwamu Tanaka, Ph.D.

Stephen P. Ficklin, Ph.D.

John Browse, Ph.D.

iii

ACKNOWLEDGEMENT

I’d like to thank the Seattle ARCS Chapter, the National Institutes of Health Protein

Biotechnology Traineeship Program, and the NSF-GRFP for monetary support. I’d like to

express my gratitude to my advisor for his time, support, and good sense of humour. Your

mentorship has changed my life for the better. I’d also like to thank my committee members for

your feedback and advice, which have allowed me to grow as a scientist. A big thanks to all of

our collaborators for sharing their time, effort and expertise to make this project possible.

Finally, I give my gratitude to my labmates, and the students, faculty, and staff associated with

the Molecular Plants Sciences program at Washington State University. I cannot imagine how

my life would be if I had not spent the past 5 years with your positive influence and

encouragement.

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THE ROLE OF KEA1 AND KEA2 TRANSPORTERS IN PLASTID ION HOMEOSTASIS

AND GENE EXPRESSION

Abstract

by Rachael Ann DeTar, Ph.D. Washington State University

May 2021

Co-Chairs: Hans-Henning Kunz, Helmut Kirchhoff

The maintenance of ion gradients across the chloroplast envelope plays a key role in the

bioenergetics of photosynthesis by moderating proton motive force and membrane dynamics.

However, this research supports the novel finding that plastid ion transporters are additionally

involved in maintaining plastid gene expression (PGE), likely by impacting the stromal buffer

conditions required for function of plastid RNA binding proteins. We discovered this by

investigating mutant Arabidopsis thaliana lines with loss-of-function of two inner envelope

membrane potassium-proton (K+/H+) antiporters KEA1 and KEA2. Simultaneous loss of both

transporters results in a unique “virescent” phenotype in which young leaves have

disproportionately lower photosynthetic efficiency, chlorophyll production, and underdeveloped

chloroplasts compared to older leaves. The goal of this research was to determine how the loss of

KEA1/2 transporters results in this peculiar chloroplast developmental phenotype. Preliminary

experiments using Total-reflection X-ray Fluorescence (TXRF) revealed that loss of KEA1/2

perturbs overall plastid ion homeostasis. An analysis of nuclear and plastome gene expression

revealed significant defects in plastid ribosomal RNA processing in kea1kea2 mutant lines. This

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likely results from altered stromal ion concentrations inhibiting the function of nuclear-encoded

chloroplast RNA binding proteins involved in plastid gene expression. This defect coincided

with decreased steady-state levels of photosynthesis-related proteins, and lower translation rates

in the stroma. We also discovered that plastid-to-nucleus retrograde signaling protein Genomes

Uncoupled 1 (GUN1) was essential to seedling survival in kea1kea2. Loss of GUN1 in the

kea1kea2 background resulted in higher expression of many nuclear-encoded photosynthesis-

associated genes which are normally suppressed in response to disruption of PGE. In summary,

ionome-induced impairment of plastid gene expression and subsequent retrograde signaling to

suppress nuclear gene expression culminates in the virescent phenotype displayed by the

kea1kea2 mutants. These findings underscore the importance of ion transporters in chloroplast

development.

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TABLE OF CONTENTS

Page

ACKNOWLEDGMENT................................................................................................................ iii

ABSTRACT ................................................................................................................................... iv

LIST OF TABLES ........................................................................................................................ vii

LIST OF FIGURES ..................................................................................................................... viii

CHAPTER

CHAPTER 1: INTRODUCTION ........................................................................................1 CHAPTER 2: CHLOROPLAST IONOME OF ARABIDOPSIS THALIANA MEASURED BY TOTAL REFLECTION X-RAY FLUORESCENCE ANALYSIS (TXRF) ...............................................................................................................................20 CHAPTER 3: PERTURBATION OF CHLOROPLAST ION TRANSPORT INFLUENCES NUCLEAR AND PLASTID GENE EXPRESSION ...............................56 CHAPTER 4: DISRUPTION OF PLASTID GENE EXPRESSION DUE TO LOSS OF KEA1/2 RESULTS IN ALTERED EXPRESSION OF PHOTOSYNTHESIS-ASSOCIATED NUCLEAR-ENCODED GENES (PHANGS) VIA GENOMES UNCOUPLED 1 (GUN1) MEDIATED RETROGRADE SIGNALING .......................126 CHAPTER 5: CONCLUSION ........................................................................................155

APPENDIX

APPENDIX A: THEORETICAL OVERVIEW OF SELECTED METHODS ..............159

APPENDIX B: MUTANT LINES AND GENOTYPING PRIMERS ............................167

vii

LIST OF TABLES

Page

Table 2.1: Literature values for the plastid ionome .......................................................................22

Table 2.2: Leaf ion concentration in A. thaliana and two iron (Fe) metabolism mutants .............29

Table 2.3: The Ionome of the WT plastid ......................................................................................35

Supp. Table 2.1: Element Analysis of NIST reference material ....................................................47

Supp. Table 2.2: TXRF measurements of standards spiked into isolated chloroplasts .................47

Table 3.1: Leaf-level concentrations of elements (µg*mg DW -1) ................................................64

Table 3.2: Differentially Expressed Genes (DEGs) .......................................................................69

Supp. Table 3.1: RNA sequencing statistics ................................................................................108

Supp. Table 4.1: RT-qPCR Primers .............................................................................................149

Supp. Table 4.1: RT-qPCR Protocol ...........................................................................................149

Appendix B: Mutant lines and genotyping primers .....................................................................167

viii

LIST OF FIGURES

Page

Fig. 1.1: Research Aims. ................................................................................................................13

Fig. 2.1: Phenotypes of iron metabolism mutants fro7 and opt3-2 ................................................27 Fig. 2.2: Isolation and evaluation of intact chloroplasts ................................................................31 Fig. 2.3: Isolation protocol yields high proportion of intact plastids .............................................32 Fig. 2.4: Preparation and measurement of chloroplasts using TX-RF analysis .............................34 Fig. 2.5: Elemental analysis of isolated plastids from WT, fro7, and opt3-2 using TXRF ...........36 Fig. 2.6: Elemental analysis of plastids from WT and kea1kea2 using TXRF ..............................38 Fig. 2.7: Model of how loss of K+/H+ exchangers KEA1/2 influences leaf plastid ionome..........42 Supp. Fig. 2.1: A sample of chloroplasts digested in HNO3 .........................................................48 Supp. Fig. 2.2: Analysis of plastids from WT, fro7, and opt3-2 norm. to chlorophyll .................49 Supp. Fig. 2.3: Analysis of plastids from WT and kea1kea2 norm. to chlorophyll ......................50 Supp. Fig. 2.4: Elemental analysis of HNO3-digested plastids......................................................51 Fig. 3.1: An overview of K transport protein in A. thaliana ..........................................................58 Fig. 3.2: kea1kea2 double mutants phenocopy many PGE mutants ..............................................62 Fig. 3.3: KCl treatment exacerbates the kea1kea2 photosynthetic phenotype, while NaCl rescues phenotype ..........................................................................................................................66 Fig. 3.4: PCA Analysis ..................................................................................................................68 Fig. 3.5: SUBA4 Multiple Marker Abundance Profiling (MMAP) tool applied to lists of Differentially Expressed Genes (DEGs) reveals many differentially expressed transcripts in kea1kea2 encode chloroplast-targeted proteins .............................................................................70 Fig. 3.6: Functional enrichment analysis reveals that significantly Differentially Expressed Genes (DEGs) in kea1kea2 compared to the WT under control conditions are often associated with specific GO terms, INTERPRO domains and PFAM families ............................72

ix

Fig. 3.7: Log2 fold-change expression of individual genes that are significantly differentially expressed in kea1kea2 compared to WT under control conditions are no longer differentially expressed when comparing kea1kea2 and WT when both are treated with NaCl .........................74 Fig. 3.8: Log2 fold change expression of genes with annotations of interest ................................76 Fig. 3.9: Average relative translation output, transcript accumulation and translation efficiency of kea1kea2 compared to WT show no substantial specific changes in plastid gene expression under NaCl stress vs control conditions. However, kea1kea2 exhibits a plastid rRNA processing defect ............................................................................................................................78 Fig. 3.10: Loss-of-function lines with plastid 16S rRNA maturation defects resemble kea1kea2 ........................................................................................................................................80 Fig. 3.11: Plastid rRNA processing defects in the kea1kea2 mutant are alleviated under NaCl stress conditions compared to control conditions. .........................................................................82 Fig. 3.12: Immunoblots reveal lower steady-state levels of plastid-encoded proteins ..................84 Fig. 3.13: Pulse-chase experiments reveal decreased translation rates in kea1kea2 .....................85 Fig. 3.14: Loss of KEA1/2 suppresses leaf variegation in var2-5 .................................................87 Fig. 3.15: Chloroplast protein import rates are similar in kea1kea2 vs WT, indicating that import of RNA binding proteins is not impaired ...........................................................................88 Fig. 3.16: Model depicting the influence of chloroplast ion homeostasis on Plastome Gene Expression (PGE) in WT plants, and in the kea1kea2 mutant under control and salt- treatment conditions .......................................................................................................................97 Supp. Fig. 3.1: Low concentrations of MgCl2 rescued kea1kea2 photosynthetic phenotype ......107 Supp. Fig. 3.2: Density plot of Fragments per Kilobase of Mapped reads (FPKM) for all replicates .....................................................................................................................................109 Supp. Fig. 3.3: Volcano Plots of RNA Sequencing Comparisons ...............................................110 Supp. Fig. 3.4: Venn Diagram of Overlapping DEGs for control and NaCl comparisons ..........111 Supp. Fig. 3.5: Replicate immunoblots ........................................................................................112 Supp. Fig. 3.6: Additional autoradiographs from the pulse-chase analyses ................................113 Supp. Fig. 3.7: Pulse-Chase in ambient light ...............................................................................114

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Fig. 4.1: A model from Martin et. al. 2016 showing opposing roles of informational light and GUN1-mediated retrograde signaling in controlling expression of the PhANGs. ......................130 Fig. 4.2: GLK1/2 transcription factors play an important signaling role in kea1kea2. ...............133 Fig. 4.3: Hypocotyl Elongation under monochromatic light for WT, kea1kea2, and phya or phyb controls ...............................................................................................................................135 Fig. 4.4: DAB stain for H2O2 production shows kea1kea2 accumulates more ROS than the WT under control conditions, but not under salt treatment .........................................................136 Fig. 4.5: Retrograde signaling via GENOMES UNCOUPLED 1 (GUN1) is triggered in response to loss of KEA1/KEA2 to downregulate Photosynthesis-Associated Nuclear- encoded Genes (PhANGs). ..........................................................................................................137 Fig. 4.6: Transcript level expression of GLK1 and a selection of downstream PhANGs in 24- day-old WT, kea1-1kea2-1, gun1-201kea1-1kea2-1, and gun1-201 ...........................................139 Fig. 4.7: A proposed model for retrograde signaling in response to loss of KEA1/2 ............................143 Supp. Fig. 4.1: Chlorophyll A, Chlorophyll B, and Chlorophyll A/B ratios in gun1, kea1kea2, and triple mutants .........................................................................................................................148 Suppl. Fig. 4.2: Theoretical maximum quantum efficiency of PSII (Fv/Fm) in 24-day-old WT, kea1-1kea2-1, gun1-201kea1-1kea2-2, and gun1-201 seedlings ........................................148 Fig. 5.1: Research Aims ...............................................................................................................155 Fig. 5.2: Model depicting consequences of KEA1/2 loss-of-function for plastid ion homeostasis, Plastome Gene Expression (PGE), and retrograde signaling .................................156 Fig. A1: Three common methods of elemental analysis..............................................................159 Fig. A2: Schematic of typical TXRF setup ......................................................................................161 Fig. A3: The three fates of light energy absorbed by PSII ..........................................................162 Fig. A4: An example of a chlorophyll fluorescence spectra and the meaning of various peaks ...........................................................................................................................................163

xi

Dedication

To Dr. Richard Martyr and Mrs. Chris Chou.

Public high school teachers can and do make a difference in their students’ lives.

You certainly made a difference in mine.

1

CHAPTER 1: INTRODUCTION

Chloroplast ion homeostasis- the intersection between plant nutrition and photosynthesis

A ubiquitous phrase in many scientific publications, grants, and dissertations is “food, fiber and

fuel”, which underscores the importance of plant research to address the growing world population.

Different models for food demand project a 58-98% increase in food production will be required

by the year 2050 to sustain the world population (Valin et al., 2014). This grim figure does not

account for the fact that many regions already suffer from food shortages and malnutrition (Lynch,

2007). However, the challenge of increasing food production to prevent mass famine has been

overcome in the past during the so-called “Green Revolution.” Two factors were primarily

responsible for this phenomenon: availability of new, high yielding varieties of staple crops

(Evenson and Gollin, 2003), and improved agronomic inputs, such as fertilizer (Lynch, 2007).

With the approaching Malthusian crisis, many researchers have pointed to the need for a second

Green Revolution (Wollenweber et al., 2005; Lynch, 2007). Just as in the latter half of the 20th

century, plant scientists could take a bipartite approach to meeting food demand: we can attempt

to improve yield by addressing how we grow plants (agronomy), and/or by improving the plants

we grow (genetics). However, our potential for yield improvement via agronomy may be limited.

Many developed nations have maxed out the potential agronomic inputs, including soil amendment

and irrigation (van Ittersum et al., 2013). Furthermore, these intensive production practices often

have negative impacts on the environment and surrounding ecosystem (reviewed in Tilman et al.,

2002). Developing nations often have more potential to increase yield by improvements in

agronomy, but for economic reasons most farmers are unwilling, or unable to risk resources to

change agronomic practices (George, 2014). Therefore, the primary avenue for increasing crop

productivity may be genetic improvement. One of the most promising and challenging proposals

2

for genetic improvement of crop yields is to manipulate photosynthesis on a fundamental level

(reviewed in (Long et al., 2015; Araus et al., 2019)). Studies suggest we will soon max out potential

yield gains by altering how plants use photoassimilates (partitioning efficiency) and how much

sunlight crops can absorb (interception efficiency (Long et al., 2015)). However, there is

considerable untapped potential in improving photosynthesis by altering how plants use the

sunlight they absorb to create photosynthates (conversion efficiency, (Long et al., 2015)). Thus, to

achieve the dramatic increases in yield needed to feed the planet’s future population of 10 billion,

plant scientists should target conversion efficiency of photosynthesis.

Photosynthesis is a complex set of reactions that we conceptualize as being divided into

light harvesting (light-dependent) reactions in the thylakoid membrane electron transport chain

(ETC) and carbon fixation (light-independent reactions in the chloroplast stroma). Both sets of

reactions have been the targets for increasing photosynthetic conversion efficiency. For example,

there have been efforts to alter the stoichiometry of the chloroplast ATP synthase rotor so that

more ATP can be produced with lower ion motive force (Pogoryelov et al., 2012). Reengineering

photosynthesis is promising in theory, but likely will lead to unforeseen complications if not

considered in the greater context of chloroplast physiology. The chloroplast is a highly complex

organelle, composed of multiple compartments with unique chemical properties. For instance, the

stroma has a pH of 7-8 (Wu and Berkowitz, 1992), whereas the thylakoid lumen has a pH of ~5

(Iwai et al., 2008) when illuminated. A complex system of ion gradients is interlinked with pH

(Höhner et al., 2016b). We are only beginning to understand that these chemical gradients have

roles beyond simply driving ATP production. Indeed, many processes required for chloroplast

function only operate under specific stromal and lumen buffer conditions. It would be

counterproductive to alter the ATP synthase dynamics without accounting for the impact on other

3

processes mediated by chloroplast ion gradients. Ion transporters and channels play a key role in

moderating the ion gradients both in the lumen and the stroma, and thus could be used to counteract

some of the side effects of re-engineering photosynthesis. Thus, the emerging field of organellar

ion transport is valuable for increasing crop yields. The focus of this work will be to untangle how

loss of chloroplast ion transporters influences photosynthetic productivity and chloroplast function

in Arabidopsis thaliana. This research will hopefully provide useful insights for engineering

photosynthetically improved plants.

A general overview of plant element needs and nutrition

Carbon (C), hydrogen (H), oxygen (O), nitrogen (N), sulfur (S), phosphorus (P), calcium (Ca),

magnesium (Mg), and potassium (K) are the most abundant elements in living tissue by mass, the

so-called macronutrients of life (Maathuis, 2009). Additionally, many organisms are also reliant

on other elements in smaller quantities known as micronutrients, including iron (Fe), manganese

(Mn), boron (B), zinc (Zn), copper (Cu), molybdenum (Mo), chlorine (Cl) and nickel (Ni)

(Marschner, 2012). Higher plants require all these elements in order to complete their life cycle,

although some plant species require additional elements, such as silicon, to survive (Epstein,

1999). Regardless, all plant species face the unique challenge of being sessile organisms that must

obtain these elements from the predetermined surrounding environment where they grow. Plants

must uptake these nutrients from the substrate and mobilize these elements to the tissues where

they are needed. Carbon and oxygen are readily available to the plant in the form of CO2 fixed

during the Calvin-Benson Cycle of photosynthesis. Hydrogen is also ubiquitous as it enters the

plant as water. These elements generally only limit plant growth when environmental conditions

impede plants’ ability to carry out oxygenic photosynthesis. However, all other essential elements

are available exclusively from the substrate on which the plant lives and are thus known as mineral

4

nutrients. Some of these nutrients are taken up as organic compounds, but generally most mineral

nutrients are taken up as soluble salts, which can be polyatomic compounds (i.e., PO43-, SO4

2-,

NO3-, NH4

+), or monoatomic ions, i.e. K+, Ca2+, etc. The fate of most polyatomic nutrients

including P, N, and S is to be assimilated into macromolecules such as nucleic and amino acids.

Transition metal mineral nutrients such as Fe, Mo, Zn, and Cu often function as cofactors in

enzymes, particularly those involved in oxidation reduction reactions. Finally, elements taken up

as soluble ions such as K, Ca, Mn, Mg, and Cl typically act as regulators of osmotic and

electrochemical potential across membranes (Marschner, 2012), although Mg and Mn also act as

cofactors in macromolecules (Waters, 2011; Alejandro et al., 2020). To some degree, cations like

K, Ca, Mg, and even Na are interchangeable for moderating electrochemical and pH gradients.

These cations also nonspecifically moderate the activity of enzymes, ribosomes, and other cellular

machinery via ionic interactions (Leigh and Jones, 1984; Maathuis, 2009; Marschner, 2012).

However, of the cations, K almost exclusively remains as a free, soluble ion within the plant (Leigh

and Wyn Jones, 1984; Marschner, 2012). This makes K the most important element in vivo for

maintaining water potential (Maathuis, 2009; Marschner, 2012; Sharma et al., 2013). Potassium

also plays a significant role in pH moderation across cellular and subcellular membranes by acting

as a counterion to H+, often via K+/H+ antiporters (Sze and Chanroj, 2018). Within the cell, the

green chloroplast has high demand for all of the afore-mentioned nutrients, as significant

bioenergetic and metabolic pathways are localized there. All mineral nutrients play some essential

role in chloroplast function and photosynthesis.

Mineral nutrients in the chloroplast and their effect on photosynthesis

The chloroplast is the highest-maintenance organelle in terms of ATP use and protein production

(Li et al., 2017). Not surprisingly, the distribution of plant mineral nutrients also reflects the large

5

investment of resources required for photosynthesis and other chloroplast processes; most Ca, K

and P imported by the roots is transported to the photosynthetic tissues (Conn and Gilliham, 2010).

In particular, transition metal and alkali earth metal nutrients have a very important role in

photosynthesis, both as structural components of photosynthetic proteins and as free ions.

Mineral nutrients and their role in photosynthetic energy transfer and electron transport.

One of the primary uses for mineral nutrients in the chloroplast is to facilitate energy transfer and

electron transport during photosynthesis. Oxygenic photosynthesis relies on a string of cofactors

with increasing reduction potential to move electrons through the ETC and drive the production of

ATP/NADPH. Most of these cofactors are transition or alkali earth metals, due to these elements’

ability to mediate redox chemistry. Moving sequentially through the ETC, the first cofactor-

containing molecule is chlorophyll, the primary plant pigment involved in photosynthetic energy

absorption and transfer. Chlorophyll carries a central magnesium (Mg2+) ion within its chlorin ring.

Depending on species and growth condition, upwards of 15% of leaf magnesium can be bound in

chlorophyll molecules (Bohn et al., 2004). Next, Manganese-calcium (Mn-Ca) clusters in the

Oxygen Evolving Complex (OEC) are required to split water and provide electrons to be excited

by harvested light energy (Blankenship). Interestingly, the OEC is also the only known plant

protein complex that requires chloride (Cl-) as a cofactor (Kawakami et al., 2009). Copper (Cu) is

the cofactor in the electron carrier plastocyanin (Aguirre and Pilon, 2016). Numerous Fe-

containing molecules, such as hemes and Fe-S clusters, are complexed into PSII, the cytochrome

B6F complex, PSI and ferredoxin to act as electron donors and acceptors. Not surprisingly, up to

80% of leaf Fe is located in the chloroplast (Kroh and Pilon, 2020). Thus, metal nutrients are

essential to photosynthesis, playing a direct role in energy harvesting and electron transfer as

protein-bound cofactors through the entire length of the ETC.

6

Soluble ionic nutrients interact with thylakoid membranes, regulating chloroplast ultrastructure

and state transitions.

In addition to being part of protein complexes, many nutrients play a key role in chloroplasts as

soluble monoatomic ions. Notably, soluble ion gradients can impact the architecture of the

thylakoids, the internal membranes of the chloroplast which house light harvesting and electron

transport machinery (Kaňa and Govindjee, 2016). The thylakoid membranes can stack vertically

into appressed structures called grana or remain as unstacked regions protruding into the stroma

known as stroma lamellae. The grana are narrow and tightly packed with PSII, whereas stroma

lamellae house bulkier proteins such as PSI (Kirchhoff, 2013). The ratio of grana to stroma

lamellae changes dynamically in response to light conditions. Mechanistically, this is

accomplished by manipulating stromal concentrations of cations which interact with negatively

charged membrane surfaces to form an Electrical Double Layer (EDL) (Kaňa and Govindjee,

2016). In the EDL, Mg and K concentrate on both surfaces of the thylakoid membrane to screen

negatively charged proteins and lipid headgroups, a phenomenon called “electrostatic screening.”

High screening due to sufficient concentrations of Mg2+ and K+ promotes thylakoid stacking into

grana because the EDL prevents repulsion from opposing thylakoid membranes (Puthiyaveetil et

al., 2017). Grana stacking occurs in low-light conditions to maximize photon capture by the light

harvesting complexes. Conversely, grana de-stack in high light conditions as the machinery needed

to repair photodamaged PSII is located in the stroma lamellae (Kirchhoff, 2013). Additionally,

high light conditions trigger Cl- ion influx which, in turn, drives thylakoid swelling (Herdean et

al., 2016a). Thylakoid swelling allows more space and higher diffusion rates in the cramped grana,

which also promotes PSII repair and speeds up the movement of plastocyanin (PC) to keep pace

7

with light harvesting (Kirchhoff et al., 2011; Herdean et al., 2016b). Thus, soluble monoatomic

elements are essential for optimizing photosynthesis via interactions with membranes.

While not directly related to thylakoid architecture, it is important to note that high

electrostatic screening via cations promotes separation of PSI/PSII, which prevents direct energy

transfer or ‘spillover’ between the photosystems. Electrostatic screening also regulates the

movement of light harvesting complexes (LHCs) between PSI and PSII to balance excitation of

the photosystems, i.e., state transitions (reviewed in Kaňa and Govindjee, 2016). This further

underscores the importance of ions in the EDL for regulating photosynthesis.

Soluble ions maintain proton motive force (PMF) across the thylakoid membrane.

Another key role of soluble ions is to regulate the voltage gradient (∆Ψ) and concentration gradient

(∆pH) components of Proton Motive Force (PMF). The individual contributions of ΔpH and ΔΨ

to PMF is called PMF partitioning. Although the magnitude of the PMF is the only determining

factor for ATP production by ATP synthase, PMF partitioning can influence the regulatory

processes of photosynthesis (Hangarter and Good, 1982; Kaňa and Govindjee, 2016). In

mitochondria, PMF is primarily stored as ∆Ψ, whereas the chloroplast maintains PMF primarily

as ∆pH. Thus, the relative contributions of the ∆Ψ and ∆pH components of PMF is altered by ion

transfer mechanisms that are independent from the ETC. It is hypothesized that ∆pH is built at the

expense of ∆Ψ through the export of K+ and Mg+ to the stroma and import of Cl- counterions to

the thylakoid lumen (Kaňa and Govindjee, 2016; Armbruster et al., 2017). The maintenance of

∆pH in the thylakoid lumen is very important for short-term photoprotection at high light

intensities. Low pH triggers nonphotochemical high energy quenching (qE), a process whereby

excess light energy absorbed by PSII is dissipated as heat (Müller et al., 2001; Ruban, 2016). Low

luminal pH also slows plastoquinol oxidation by cytochrome b6f to limit the rate of electron

8

transport (Tikhonov, 2013; Armbruster et al., 2017). This type of regulation, referred to as

“photosynthetic control”, prevents PSI photodamage (Colombo et al., 2016). All in all, increased

ΔpH protects plants from photodamage in high light, but can reduce flux through the

Photosynthetic Electron Transport Chain (ETC) upon transition to low light. Thus, the moderation

of ΔpH is especially important when plants grow in fluctuating light conditions, where NPQ must

be rapidly induced and dissipated to optimize photosynthesis and minimize photodamage. Ion

transporters and channels help the chloroplast maintain and partition the PMF in fluctuating light

conditions (Armbruster et al., 2017).

Ions maintain buffer conditions required for enzyme and ribosome activity in the chloroplast.

The chloroplast stroma and thylakoid lumen maintain a unique chemical environment. For

example, under illumination the cytosol has a pH of 7, the stroma a pH of 8, and the lumen a pH

of 6 (Höhner et al., 2016a). Differing concentrations of soluble ions are also maintained in these

compartments. Most enzymes have evolved to operate under specific conditions within these

compartments, which often change with light intensity. Thus, stromal and luminal pH and ion

concentrations are moderated in these compartments for two reasons: 1) To link enzyme activity

to the light cycle. 2) To keep enzymes and other machinery running in spite of the light cycle. A

good example is the light-induced activation of the Calvin-Benson-Bassham (CBB) cycle. The

rate-limiting enzyme RuBP carboxylase (Rubisco), is activated by stromal alkalization (Mott and

Berry, 1986). In turn, the stromal alkalization is codependent upon the influx of soluble cations

such as K+, Ca2+, and Mg2+ to counterbalance H+ efflux in the light (Ishijima et al., 2003;

Armbruster et al., 2017). These influxes of stromal calcium and magnesium also directly regulate

enzymes of the CBB cycle, including Rubisco (reviewed in Pottosin and Shabala 2016). For

example, high Mg2+ concentrations stabilize Rubisco activase, which in turn activates Rubisco by

9

removing inhibitors bound to the enzyme (Hazra et al., 2015). Intriguingly, this effect seems to

dominate the direct inhibition of Rubsico by Mg2+ binding to the active site (Liang et al., 2008).

Beyond enzymes involved in bioenergetic processes, proper plastid gene expression is also

reliant on stromal ion and pH concentrations based on in-vitro experiments (Bhaya and Jagendorf,

1984; Horlitz and Klaff, 2000; Draper et al., 2005; McDermott et al., 2018; Gawroński et al.,

2020). Sensitivity to stromal buffer conditions may come into play on the posttranscriptional and

translational levels. A major step in plastome gene expression is post-transcriptional RNA

processing, which is mediated by stroma-targeted nuclear-encoded proteins, i.e., pentatricopeptide

repeat proteins (PPRs; Tillich et al., 2010; Zoschke et al., 2011). In-vitro, sodium chloride (NaCl)

alters chloroplast RNA secondary structure and thus impairs the binding of PPRs to target proteins

(McDermott et al., 2018). While it has not yet been investigated experimentally, it is possible the

binding ability of other plastid-localized RNA processing proteins such as endo- and exonucleases

(Stoppel and Meurer, 2011) could also be impaired by extreme RNA secondary structure induced

by high ion content. The degree to which posttranscriptional processing of mRNA influences the

likelihood of translation into a functional polypeptide is still an open question in the field. Yet,

many mutants with plastid RNA processing defects do exhibit lower translation rates (Kleinknecht

et al., 2014), thus perturbation of RNA processing via altered stromal ion content could impair

overall plastid proteostasis.

Furthermore, plastid gene expression has been shown to be directly inhibited on the

translational level in vitro. Previous studies have shown bacterial-type 70s ribosomes, like those

in plastids, require sufficient - but not excessive - concentrations of cations such as Mg2+, and K+

for assembly and function (Bhaya and Jagendorf, 1984; Horlitz and Klaff, 2000; Blaha et al., 2002;

Hirokawa et al., 2002; Konevega et al., 2004; Petrov et al., 2012; Nierhaus, 2014). In theory,

10

altered stromal levels of these cations could therefore impair translation rates of plastome-encoded

transcripts in-vivo, although this has never been tested. Clearly ions play a key role in the overall

functioning of the chloroplast, yet the relationship between plastid gene expression and stromal

ion homeostasis remains an open area for research.

Phenotypes of loss-of-function mutants for chloroplast ion transporters

Significant progress identifying and characterizing chloroplast ion transport proteins has been

made in recent years by using Arabidopsis thaliana loss-of-function lines (for review see Höhner

et al., 2016a; Szabo and Spetea, 2017). Chloroplast ion transport protein loss-of-function mutants

exhibit phenotypes corresponding to the localization of the transporter (envelope vs. thylakoid)

and the ion(s) transported. Interestingly, loss of thylakoid ion transport generally has a specific

effect on photosynthetic energetics, with little overall phenotypic impact. Conversely, loss of

chloroplast envelope channels and transporters seem to trigger pleiotropic effects on plant

phenotype.

Thylakoid membrane transport mechanisms

Thylakoid transporters often play an important role in photosynthetic regulation. For example, null

mutants for thylakoid K+/H+ antiporter KEA3 have increased PMF partitioning towards ΔpH ,

which leads to increased high-energy nonphotochemical quenching (qE) at low light intensities

(Armbruster et al., 2014). KEA3 moderates H+ efflux from the lumen to relax qE and divert energy

back to photochemistry. Conversely, loss-of-function lines for voltage-gated Cl- channel

VCCN1/BEST1 have decreased ΔpH, and thus cannot reach WT-levels of qE in fluctuating high-

light conditions (Duan et al., 2016; Herdean et al., 2016b). VCCN1/BEST1 is hypothesized to

import Cl- into the lumen in the light to counterbalance H+ influx via the ETC, thus loss of this

transporter increases the ΔΨ portion of PMF (Duan et al., 2016). Another thylakoid membrane Cl-

11

channel CLCE is theorized to export Cl- from the thylakoid lumen during the light to dark transfer.

Knockout lines for CLCE also exhibit altered PMF partitioning but these effects are only dramatic

in the dark (Herdean et al., 2016a). Loss of KEA3, VCCN1/BEST1, and CLCE affects thylakoid

structure, but otherwise does not result in an obvious visual phenotype (i.e., reduced biomass or

chlorosis) under laboratory light conditions. Fluctuating light treatment causes kea3 mutants to

acquire less biomass than WT plants (Armbruster et al., 2016). Thus, thylakoid transporters are

primarily important for moderating the bioenergetics of photosynthesis under natural light

conditions. The exception is for loss-of-function of PAM71, a thylakoid protein that likely

transports Mn/Ca (Schneider et al., 2016). Plants missing PAM71 do have an obvious phenotype,

specifically chlorosis and reduced biomass, but this is likely because PAM71 transports Mn

cofactors needed for the OEC on the luminal side of the thylakoid membrane.

Plastid Envelope transport mechanisms

In contrast to thylakoid channels and carriers, loss-of-function mutants for plastid envelope

transport proteins typically have dramatic visual phenotypes regardless of growth light condition.

For instance, loss of K+ Efflux Antiporters (KEA) results in reduced photosynthetic light

harvesting efficiency, reduced biomass, and chlorotic young leaves (Kunz et al., 2014). Decreased

chlorophyll content and low photosynthetic efficiency specifically in young leaves is characteristic

of ‘virescent’ mutants with chloroplast development defects. Mutants for chloroplast envelope Mn

transporter CMT1 exhibit overall chlorosis (Eisenhut et al., 2018) and loss of chloroplast envelope

Cl- channels MSL2/MSL3 result in leaf variegation (Haswell and Meyerowitz, 2006). The shape

and size of chloroplasts in all these lines also differs significantly from WT. In msl2msl3, the

perturbation of chloroplast structure and size is posited to be due to osmotic (rather than

developmental) effects, as even non-green plastids in msl2msl3 are swollen and oversized (Haswell

12

and Meyerowitz, 2006; Veley et al., 2012). In contrast, kea1kea2 has some large swollen

chloroplasts that would be expected due to osmotic imbalance, but additionally has an unusually

high number of small, etiolated plastids in leaf tissue (Aranda-Sicilia et al., 2016). Furthermore,

kea1kea2 lines accumulate lower steady-state levels of chloroplast proteins, particularly in young

leaves (Aranda-Sicilia et al., 2016). The cmt1 line also exhibited a heterogenous population of

chloroplasts, and reduced levels of certain plastid proteins, although experiments did not

differentiate between leaf ages (Eisenhut et al., 2018). Taken together, the virescent phenotype,

reduced levels of chloroplast proteins, and aberrant chloroplast development in these lines

indicates that envelope ion transporters play an important role in plastid development. Yet, it is

unknown how cation homeostasis mechanistically impacts plastid development. Given the reliance

of plastid RNA binding proteins and ribosomes on proper ion concentrations in vitro, one

promising hypothesis is that plastid envelope transporters maintain optimal levels of cations for

plastid gene expression. The overall goal of this research is to test this hypothesis, and thus better

understand the pleiotropic role of plastid envelope ion transporters. This goal was broken down

into 3 aims: 1) Quantify the effect of KEA1/2 loss-of-function on chloroplast ion concentrations;

2) Determine if plastid gene expression is altered by loss of KEA1/2; 3) Characterize how plants

sense and respond to loss of chloroplast ion homeostasis.

13

Fig. 1: Research aims

14

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CHAPTER 2: CHLOROPLAST IONOME OF ARABIDOPSIS THALIANA MEASURED BY TOTAL REFLECTION X-RAY FLUORESCENCE ANALYSIS (TXRF).

This chapter is written and formatted in the style of a Plant Methods journal article. We anticipate submitting this work for publication soon with coauthors Drs. Ricarda Höhner, Bettina Bölter and Hans-Henning Kunz.

ABSTRACT

Background: The regulation of ion flux, i.e., ion homeostasis, is crucial for bioenergetic organelles such as the chloroplast. Plastids harness ion gradients to build and manipulate proton motive force (PMF), activate enzymes in the Calvin Cycle, and regulate other metabolic and developmental processes. Additionally, many plastid proteins require ions as cofactors. In recent years, there has been a surge in publications characterizing newly discovered plastid ion transport proteins in Arabidopsis thaliana. However, the ability to measure how these transport proteins influence the concentration of various elements within the chloroplast remains challenging due to various technical problems, particularly the difficulty of obtaining large volumes of intact A. thaliana chloroplasts needed for conventional ionomics protocols.

Results: We utilize a recent technology in the field of ionomics, Total Reflection X-Ray Fluorescence (TXRF) analysis to overcome many of the obstacles associated with chloroplast ionomics. We optimize the TXRF sample preparation, measurement, and analysis to accommodate small volumes of highly dense chloroplast preparations. Additionally, we altered the standard A. thaliana chloroplast preparation method to obtain many intact plastids from A. thaliana mutants. We were able to use our method to measure the ionome of plastids isolated from iron (Fe) metabolism mutants opt3-2 and fro7, and plastid K+/H+ antiporter loss-of-function mutant kea1kea2. While our method did not reproduce the low-Fe phenotype previously shown for fro7 plastids, we were able to quantify Fe over-accumulation in opt3. Additionally, our approach yielded promising preliminary data for the plastid ionome in kea1kea2. Our preliminary data indicate KEA1/2 loss-of-function causes mutant chloroplasts to over-accumulate K+ and other elements, thus supporting the hypothesis that the KEA1/2 transporters act as a valve for K+ from the stroma.

Conclusions: Our method will enable researchers to directly link phenotypes of chloroplast ion transporter mutants with alterations in the chloroplast ionome. This allows direct conclusions to be made about the activity and function of chloroplast ion transporters, and to what degree their loss or gain of function impacts plastid ion homeostasis. Furthermore, our method can be modified to directly measure the chloroplast ionome of other A. thaliana mutants of interest.

Keywords: ionomics, plastid, photosynthesis, iron, potassium.

21

BACKGROUND

Ion and proton gradients across membranes act as a battery for the cell, providing a source of

potential energy which can be rapidly harnessed to energize numerous chemical and physical

processes. The chloroplast is a site where this cellular battery is charged and discharged during the

process of photosynthetic electron transfer. Light from the sun is converted into chemical and

electrical potential via H+ transfer across the thylakoid membrane. This potential energy, called

Proton Motive Force (PMF), is then leveraged to generate ATP, the energy currency of the cell.

PMF can also be driven by soluble ion gradients, which can be generated by protein-facilitated

electroneutral exchange of protons for cations across the thylakoid membrane (Sze and Chanroj,

2018). Thus, mechanisms by which photosynthetic organisms regulate PMF and chloroplast ion

gradients in general have received great attention in recent years. Many research groups have used

the genetic resources available for model plant Arabidopsis thaliana to study the physiological

significance of ion transport proteins including channels and transporters (reviewed in Höhner et

al., 2016a; Pottosin and Shabala, 2016; Armbruster et al., 2017). Several independent studies have

implicated these transporters in maintaining different aspects of photosynthesis and plastid

function, including moderation of Non-Photochemical Quenching (Armbruster et al., 2016; Duan

et al., 2016; Herdean et al., 2016), light harvesting capacity via PSII (Fv/Fm; Kunz et al., 2014;

Eisenhut et al., 2018), and plastid development (Aranda-Sicilia et al., 2016). Yet despite the

thorough characterization of the downstream effects of loss of ion transport mechanisms in the

chloroplast, quantification of the direct effects on the plastid ionome remains rare. This disparity

results from 1) the difficulty of isolating intact A. thaliana chloroplasts; 2) the large volumes

required by conventional elemental analysis platforms; and 3) the necessity for sample digestion

by conventional elemental analysis platforms. To overcome these limitations and resolve the A.

22

thaliana chloroplast ionome, we use an updated chloroplast isolation protocol to obtain intact

chloroplasts, then measure the organelles using Total-Reflection X-ray Fluorescence (TXRF)

analysis.

Table 2.1: Literature values for the plastid ionome

Element Conc. Unit Species Method Source

Sodium

46.2 4.0 39.1 33.6 29.2 3.1

ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl

Spinach Spinach Spinach Sugarbeet Pea Pea

AAS AAS flame photometry flame photometry flame photometry flame photometry

Robinson et al 1983 Schroppel-Meier and Kaiser 1988 Robinson and Downton 1984 Robinson and Downton 1984 Robinson and Downton 1984 Nobel 1969

Magnesium 0.25 10.9 4.7

ng*10-6 plas. ng*µg-1 Chl ng*µg-1 Chl

A. thaliana Spinach Pea

ICP-MAS AAS AAS

Sun et al. 2017 Schroppel-Meier and Kaiser 1988 Nobel 1969

Chlorine

75.2 0.9 52.3 81.8 64.7 99.2 36.7

ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl

Spinach Spinach Spinach Spinach Sugarbeet Pea Pea

silver titration AAS silver titration silver titration silver titration silver titration silver titration

Robinson et al 1983 Schroppel-Meier and Kaiser 1988 Demmig and Gimmler 1983 Robinson and Downton 1984 Robinson and Downton 1984 Robinson and Downton 1984 Nobel 1969

Potassium

169.5 175.5 151.1 181.4 135.7 49.1 47.3

ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl ng*µg-1 Chl

Spinach Spinach Spinach Spinach Sugarbeet Pea Pea

AAS AAS flame photometry flame photometry flame photometry flame photometry flame photometry

Robinson et al 1983 Schroppel-Meier and Kaiser 1988 Demmig and Gimmler 1983 Robinson and Downton 1984 Robinson and Downton 1984 Robinson and Downton 1984 Nobel 1969

Calcium 8.2 ng*µg-1 Chl Pea AAS Nobel 1969

Manganese 29 1

ng*10-6 plas. ng*10-6 plas.

A. thaliana A. thaliana

ICP-MS ICP-QQQ-MS

Divol et al. 2013 Eisenhut et al. 2018

Iron 18.1 5.0 3.0

ng*10-6 plas. ng*10-6 plas. ng*µg-1 Chl

A. thaliana A. thaliana A. thaliana

ICP-MS ICP-QQQ-MS ICP-MS

Divol et al. 2013 Eisenhut et al. 2018 Jeong et al. 2008

Nickel 5.0 µg*g-1 DW rice ICP-MS, ICP-ES Li et al. 2020

Copper 1.2 ng*µg-1 Chl A. thaliana ICP-AES Seigneurin-Berny et al. 2006

Zinc 1.0 ng*µg-1 Chl A. thaliana ICP-AES Seigneurin-Berny et al. 2006

23

For elemental analysis and biochemical characterization of isolated chloroplasts, researchers

originally used spinach (Spinacia oleracea) or pea (Pisum sativum) as these species have large,

robust chloroplasts that are easy to isolate (Table 2.1; Nobel, 1969; Demmig and Gimmler, 1983;

Robinson and Downton, 1984; Schröppel-Meier and Kaiser, 1988). However, A. thaliana has

become the model organism of choice for the plant science community due to the abundance of

forward and reverse genetic tools available. Yet, isolation of intact A. thaliana plastids is

challenging as separation from lysed plastids involves centrifugation on a percoll density gradient

or pelleting after application to a percoll layer (Walker et al., 1987; Aronsson and Jarvis, 2002;

Kley et al., 2010). The osmotic status of the plastids has a strong influence on the portion of intact

plastids in any given isolation. Thus, for proof of concept we isolated and measured the ionome of

WT chloroplasts, and chloroplasts from mutant lines fro7 and opt3, which do not have impaired

chloroplast turgor, size, or structure, but do have published changes in chloroplast or leaf iron (Fe)

content respectively.

The fro7 mutant is missing a chloroplast-targeted Fe(III) chelate reductase, and

consequently has slightly lower levels of Fe in the chloroplast, as Fe(III) reduction to Fe(II)

promotes Fe uptake across the chloroplast envelope (Bughio et al., 1997; Jeong et al., 2008).

Conversely, the opt3-2 mutant lacks a phloem-targeted Fe transporter responsible for

remobilization of Fe from shoots to roots. As a result, this mutant hyperaccumulates iron in leaf

tissues (Stacey et al., 2008; Zhai et al., 2014). It is assumed that upwards of 80% of Fe is stored in

the chloroplast (Terry and Low, 1982). Therefore, it is likely that some of the excess iron in opt3-

2 is stored in the plastid and could be quantified with our ionomics method.

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We begin by isolating and measuring Col-0 (WT), fro7, and opt3-2 chloroplasts, whose

element concentrations can be normalized to total chlorophyll levels. Then we apply our protocol

to chlorotic, osmotically compromised plastids from K+/H+ antiporter mutant kea1kea2,

normalizing to chloroplast count. The kea1kea2 mutant suffers from loss-of-function of two

chloroplast envelope-localized potassium (K) Efflux Antiporters KEA1 and KEA2 (Kunz et al.,

2014). This mutant has been shown to have increased levels of potassium (K) in leaf tissue (Höhner

et al., 2016b; Höhner et al., 2019). We hypothesize that this excess K likely accumulates in the

plastid and impairs many aspects of plastid function and development. However, the plastid

ionome of kea1kea2 has yet to be characterized. The ionome of kea1kea2 plastids will present a

unique methodological challenge. As kea1kea2 plants are chlorotic, we cannot normalize

elemental concentrations to total chlorophyll (Kunz et al., 2014). Furthermore, kea1kea2 mutants

have been shown to have chloroplasts which vary dramatically in ultrastructure and developmental

stage compared to those isolated from WT plants of an equivalent age (Kunz et al., 2014; Aranda-

Sicilia et al., 2016). To combat these problems, we will apply our chloroplast isolation and TXRF

method to kea1kea2 chloroplasts and normalize element concentrations to chloroplast count. Our

hope is to be able to finally connect the chloroplast ionome to plastid development, which has been

an open question in the field for several years (Sze and Chanroj, 2018).

In our hands, plastid isolation from 5 g of WT A. thaliana leaves typically results in a small

volume of intact chloroplasts (100-300 µL of chloroplasts with less than 2000 ug*mL-1 total

chlorophyll). However, the most common multi-element analysis platforms involve oxidative

digestion of sample in large volumes of nitric acid, followed by nebulization into an Inductively

Coupled Plasma (ICP) torch to atomize elements (Wilschefski and Baxter, 2019). This process

requires a high mass of sample. Even most micro-scaled digestion methods require an initial input

25

of 1-20 mg of dry material to produce a final sample volume of about 5 mL (Hansen et al., 2009).

While some groups have been successful applying methods such as ICP Mass Spectrometry (ICP-

MS) and ICP Absorbance Emission Spectroscopy (ICP-AES) to A. thaliana chloroplasts,

presented data is limited to selected transition metal elements for which instrument sensitivity is

high (Jeong et al., 2008; Eisenhut et al., 2018). To date, no method has been developed to measure

a range of elements from a small sample of A. thaliana chloroplasts due to the limitations of ICP-

based methods. In particular, previous attempts at characterizing the ionome of A. thaliana

chloroplasts have neglected alkali earth metals like potassium (K) and calcium (Ca), which have

important physiological roles in the chloroplast (Hochmal et al., 2015; Höhner et al., 2016a;

Pottosin and Shabala, 2016). Therefore, we developed a method to measure multiple biologically

relevant elements in isolated chloroplasts using an alternate technology- Total Reflection X-ray

Fluorescence (TXRF).

TXRF uses X-rays projected onto a thin layer of sample to excite electrons from an inner

orbital of the atom. The elements are identified based on the wavelength of fluorescence released

when an electron moves to fill the vacant orbital and return the atom to ground state. Depending

on the atomic mass of the element in question, an electron can move from either a K orbital or an

L orbital to fill the empty position, corresponding to ‘K’ or ‘L’ lines on a photon energy spectrum.

The intensity of these lines can then be used to quantify a particular element in relation to a known

internal standard (Klockenkamper and Von Bohlen, 2015). This method requires only small

sample amounts and can be used on a range of different sample types- powdered solids, liquids,

suspensions, slurries or even slices of solid material (Bohlen and Reinhold, 2015). This means that

unlike for ICP-based methods, oxidation and digestion with nitric acid (HNO3) is not always

necessary. Hence, liquid samples can remain highly concentrated, allowing for quantification of

26

low-abundance elements. Indeed, this technique has already been used for trace element analysis

in Arabidopsis leaf tissue (Höhner et al., 2016b). Here, we present a method to use TXRF

instrumentation to measure elements in isolated chloroplasts from the model plant A. thaliana.

RESULTS

The visual and molecular phenotypes of fro7 and opt3 make them ideal models for examining the

chloroplast ionome.

As proof of concept, we initially used FRO7 loss-of-function mutant fro7 (Jeong et al., 2008) and

OPT3 knock-down mutant opt3-2 (Stacey et al., 2008) in addition to Col-0 (WT) A. thaliana for

chloroplast isolation and measurement. Although they have perturbed Fe metabolism, these two

mutants are not chlorotic or pale like some other mutants with perturbed ion homeostasis (Fig.

2.1A). Our measurements of chlorophyll content support this observation, as the two mutants do

not exhibit significantly altered levels of chlorophyll a or chlorophyll b compared to the WT (Fig.

2.1C). However, opt3-2 plants generally have reduced biomass along with some necrotic patches,

indicating that the perturbation of iron signaling and transport has a net effect on growth (Fig.

2.1A). As iron plays a key role in many photosynthetic proteins, we investigated photosynthetic

efficiency using Pulse-Amplitude Moderated (PAM) chlorophyll fluorescence (Fig. 2.1B, D-F).

Measured photosynthetic parameters include theoretical maximum yield of PSII (Fv/Fm, Fig. 1B,

D), quantum yield of PSII (ΦPSII, Fig. 2.1E), quantum yield of non-photochemical quenching

(ΦNPQ, Fig. 1F), and quantum yield of non-regulated non-photochemical quenching (ΦNO, Fig.

2.1G) as described in Klughammer and Schreiber, 2008. As has been shown in previous works

(McInturf et al., 2020) opt3-2 did not exhibit significant differences in the afore-mentioned

parameters compared to WT. While fro7 has been shown previously to have low ΦPSII while

27

growing on ½ MS media, our soil-grown fro7 had WT levels of ΦPSII and all other chlorophyll

fluorescence parameters. This indicates fro7 and opt3-2 lines are as photosynthetically efficient as

Fig. 2.1: Phenotypes of iron metabolism mutants fro7 and opt3-2. A RGB image of 4-week-old WT, fro7, and opt3-2 plants. B False-color image of Fv/Fm

in WT, fro7 and opt3-2. C Mean chlorophyll a (white bars), chlorophyll b (dark gray bars) and total chlorophyll (light grays bars) in leaf tissue from the three genotypes (+/- SD, n = 5). Bar graphs of photosynthetic parameters; D Fv/Fm, E ΦII, F ΦNPQ, and G Φ NO (+/- SD, n = 6). For all bar graphs, asterisks indicate statistically different means between bracketed bars (p -value < 0.05).

28

the WT, and thus will likely yield comparable chloroplasts. Since leaf-level chlorophyll is not

altered in any of the mutant lines, we can use chlorophyll levels as a robust parameter to normalize

our ionomics data.

Leaf-level ionomics reveals altered ionome for fro7, opt3-2

Initially we characterized the leaf level ionome so we could interpret how well leaf-level

alterations in ion concentrations translate to alteration of the plastid ionome. While TXRF has been

used for ionomics analysis of leaf tissue before (Bohlen and Reinhold, 2015; Höhner et al., 2016b),

we double-checked the accuracy of our instrument by doing elemental analysis of National

Institute of Standards and Technology (NIST) plant reference material (Supp. Table 2.1). Based

on these analyses, we concluded that our instrument could accurately measure concentrations of

many elements commonly found in leaves. We measured light elements from sodium (Na) to

calcium (Ca) by excitation with the tungsten L-line (W-L) beam, and heavier elements from

manganese (Mn) to strontium (Sr) with the molybdenum K-line (Mo-K) beam. Then, we calculated

recovery relative to the known values provided by the NIST, where recovery = 100 ∗

. The concentrations of most elements we measured were within 10% of the

certified value (recovery of 90-110%), indicating our instrument is sufficiently accurate to quantify

many biologically relevant elements. Elements with values that were very inaccurate (i.e., sodium,

magnesium, aluminum, and chlorine) were not presented for succeeding samples.

To analyze the leaf ionome of A. thaliana WT, fro7 and opt3-2 lines, we used whole

rosettes from 3–4 week old plants grown on soil as shown in Fig 2.1A. Subsequently, we dried

rosettes, digested them in nitric acid (HNO3), diluted the digests, then measured ion content

normalized to tissue dry weight as described in (Höhner et al., 2016b). In general, the ionome of

29

all three genotypes was very similar (Table 2.2). However, the opt3-2 mutant accumulated about

10-fold more iron (Fe) and 2-fold more manganese (Mn) than the WT or fro7. This high Fe/Mn

phenotype reproduced previous results shown for opt3-2 (Stacey et al., 2008; Zhai et al., 2014).

Opt3-2 also was the only line with reliable above-background levels of zinc (Zn) in leaf tissue.

The fro7 line did not display a leaf-level Fe phenotype, yet intriguingly accumulated

significantly higher concentrations of potassium (K) compared to opt3-2. We then moved towards

investigating if these leaf-level alterations impacted plastid ion levels. However, prior to

measuring the plastid ionome, we had to ensure we could isolate intact and comparable plastids

Table 2.2: Leaf ion concentration in WT A. thaliana and two iron (Fe) metabolism mutants.

Element

WT fro7 opt3-2

mg* g-1 DW

Std. Dev mg* g-1

DW Std. Dev

mg* g-1 DW

Std. Dev

Potassium 39.254 4.511 44.854 7.823 33.925 3.403

Calcium 41.547 4.226 46.784 7.798 43.691 4.350

Manganese 0.030 0.006 0.0308 0.004 0.0636 0.0118

Iron 0.077 0.011 0.083 0.015 0.604 0.097

Nickel Below

Backgr. N/A

Below Backgr.

N/A Below

Backgr. N/A

Copper 0.008 0.001 0.008 0.001 0.008 0.001

Zinc Below

Backgr. N/A

Below Backgr.

N/A 0.171 0.079

Mean ion concentration in mg per g dry weight (mg * g-1 DW, n = 7) as measured using Total Reflectance X-RAY Fluorescence (TXRF) on nitric acid digested rosettes, and standard deviation of the mean (Std. Dev). Bolded values indicate mean concentration differs significantly from WT concentration (p-value < 0.05). Underlined values indicate mean concentration differs significantly from other mutant line (p-value < 0.05).

30

from all genotypes of interest. This point was particularly important as elements typically present

as soluble ions (i.e., Ca, K) could readily leak from broken chloroplasts during the isolation

process, thus causing underestimation of element concentrations (Robinson and Downton, 1984).

Chloroplast isolation protocol for A. thaliana yields intact chloroplasts

To obtain intact chloroplasts from a variety of lines, we modified the chloroplast isolation protocol

from Aronsson and Jarvis 2002 (Fig. 2.2). Notably, we altered the buffers used in the protocol to

avoid exogenous application of elements of interest- e.g., we adjusted the pH of buffers with NaOH

rather than KOH. We also ran isolated plastids on a Percoll density gradient to separate intact

plastids from broken plastids and debris. The dense, intact fraction of plastids from the Percoll

gradient were viewed with phase-contrast microscopy to determine the yield of intact plastids

(Schulz et al., 2004). Chlorophyll content of the isolated plastids was also measured as described

previously (Porra et al., 1989). Observation of isolated chloroplasts with phase-contrast

microscopy revealed that chloroplasts from WT, fro7, opt3-2, kea1-1kea2-1 and kea1-2kea2-2

were all 70-80% intact (Fig. 2.3B). This indicates that soluble ion leakage due to plastid lysis

would be similar across genotypes. Therefore, differences in soluble ion concentrations between

genotypes would result exclusively from altered activity or abundance of ion transport proteins.

Another important consideration for comparing chloroplasts isolated from different

mutants was that the isolation procedure might select for different subpopulations of chloroplasts

for each mutant. For example, a chloroplast preparation from a mutant might be enriched in

plastids at a specific developmental stage (i.e., proplastids vs. developed chloroplasts), or plastids

from a specific cell type (i.e., mesophyll cell chloroplasts vs. bundle sheath). While this concern

could not be completely allayed, we calculated the total chlorophyll content per 1 million plastids

31

Fig. 2.2: Isolation and evaluation of intact chloroplasts. Red arrows on images indicate intact chloroplasts. Black arrows on images indicate broken chloroplasts.

32

for each chloroplast isolation (Fig. 2.3C). Presumably, extreme differences in chloroplast

metabolism, function, or age would be reflected in the chlorophyll content for each chloroplast.

While the total chlorophyll per million isolated chloroplasts did vary considerably, no genotype

yielded chloroplasts with a statistically significantly different chlorophyll to chloroplast ratio.

From this we conclude isolated chloroplasts from different mutants are likely somewhat similar.

Fig 2.3: Isolation protocol yields high proportion of intact plastids. A Phase contrast image of isolated plastids from (TOP) WT, fro7 and opt3-2 genotypes and (BOTTOM) WT, kea1-1kea2-1, and kea1-2kea2-2. Scale bar shows 25 µm. Broken chloroplasts are indicated by red arrows. B Mean percent yield of intact isolated plastids for each afore-mentioned genotype (+/- SD, n = 3-11). Means were not statistically significantly different from each other (p > 0.05). C Mean chlorophyll content per million isolated chloroplasts for the 5 genotypes (+/- SD, n = 3-11). Means were not statistically significantly different (p > 0.05).

33

Sample preparation for TXRF results in a flat, homogenous sample

One of the major hurdles in the use of biological material for fluorescence-based elemental analysis

is that elements in biological samples are usually embedded in a complex matrix of

macromolecules which can absorb or otherwise alter the fluorescence from the elements therein.

For most ICP applications, complete digestion of sample matrix with HNO3 negates the issue of

matrix effects by converting all elements into soluble salts. This is not a necessity for TXRF, which

overcomes this limitation by measuring an ultrathin film of sample where matrix effects are

essentially non-existent. However, this method requires the sample to be spread in a thin,

homogenous layer on the carrier. This can be accomplished in several ways; 1) Sample digestion

with HNO3 as done for ICP-based methods 2) Dilution of sample to limit thickness of viscous

samples 3) The addition of chemical agents which promote sample homogeneity.

For isolated chloroplasts, digestion with nitric acid was not an option, as a chemical

reaction between chloroplasts, buffer and nitric acid produced large quantities of gas which

resulted in bubbling on the TXRF carrier (Supp. Fig. 2.1). Hence, we had to use an alternative

method to obtain a thin layer of sample. We dilute the chloroplasts with polyvinyl alcohol (PVA)

prior to spotting on a sample carrier. This approach was successful in creating a thin film of sample

on the carrier once dried (Fig. 2.4).

We then tested our method to ensure accurate element quantification by spiking known

concentrations of reference elements into isolated chloroplasts and measuring recovery (Supp.

Table 2.2). We used vanadium (V) for measurement of light elements with a tungsten L-line (W-

L) beam, and rubidium (Rb) for measurement of heavier transition metals with the molybdenum

K-line (Mo-K) beam. The percent recovery for measured concentrations of both spiked standards

34

was 88-109% of the expected concentration. This indicated that our sample preparation method

did not create an excessively thick or heterogeneous film. Thus, we measured biologically relevant

elements in WT chloroplasts. As for the leaf tissue samples (Supp. Table 2.1, Table 2.2), we

measured light elements starting from phosphorus (K) via excitation with the W-L beam, and

Fig. 2.4: Preparation and measurement of chloroplasts using TX-RF analysis. Spectra show intensity of fluorescence in counts per second (Y-axis) emitted by elements at a characteristic energy (x-axis) after excitation by W (left spectra) or Mo X-ray tube. Black arrows indicate elements tagged for deconvolution and quantified in isolated plasmids. Red arrows indicate elements used for internal standard.

35

heavier elements from manganese (Mn) to strontium (Sr) with the Mo-K beam (Fig. 2.4). Table

2.3 shows concentrations of biologically relevant elements which were robustly quantified using

TXRF. Concentrations for elements which were not detectable, did not reliably accumulate above

background levels, or are not biologically relevant are not shown. Many of the elements displayed

a large standard deviation from the mean, particularly when normalized to chloroplast count.

However, WT chloroplast concentrations of Mn, Fe Cu, and Zn were very similar to individual

element measurements from other groups when normalizing to chlorophyll or to chloroplast count

(Table 2.3, Table 2.1). The other elements we present, including K and Ca, have never been

measured in isolated A. thaliana

chloroplasts. Intriguingly, our value

for K is ~10-fold lower than

concentrations measured in pea, and

~40-fold lower than concentrations

in spinach, and sugar beet when

normalizing to chlorophyll content

(Table 2.1). This trend merits

further investigation, ideally by

doing side-by-side isolation and

measurement of chloroplasts isolated from Arabidopsis, pea, and spinach.

TXRF measurement and analysis revealed opt3-2 mutants sequester surplus Fe in leaf plastids.

With our newly established method, we went on to measure the afore-mentioned elements in

plastids isolated from mutant lines fro7 and opt3-2. We normalized element concentrations to both

Table 2.3: The Ionome of the WT plastid

Element ng*µg-1 Chl

Std. Dev

ng*106 Plas.

Std. Dev

Potassium 3.588 3.384 14.234 20.147 Calcium 1.571 1.404 6.046 9.465

*Vanadium 0.000 0.000 0.000 0.000 Manganese 0.211 0.150 0.375 0.133

Iron 3.897 1.673 11.237 7.420 Nickel 0.024 0.030 0.046 0.035

Copper 0.923 0.680 3.682 3.262 Zinc 0.245 0.138 0.786 1.035

*Rubidium 0.000 0.000 0.000 0.000

Mean element concentration in isolated plastids normalized to total chlorophyll (ng*µg Chl-1, n = 5) or chloroplast count (ng*106 chloroplasts-1, n = 4), and standard deviation of the mean (Std. Dev). * denotes elements chosen to use as standards in Supp. Table 2.2.

36

Fig. 2.5: Elemental analysis of isolated plastids from WT, fro7, and opt3-2 using TXRF. Mean concentration of A potassium (K), B calcium (Ca), C manganese (Mn), D iron (Fe), E nickel (Ni), F copper (Cu), and G zinc (Zn) normalized to total chlorophyll (+/- SD, n = 4). Points represent values from individual samples. No statistically significant differences between genotypes were detected (p -value < 0.05). Points with corresponding color represent replicates isolated on the same day.

37

total chlorophyll (Fig. 2.5) and to chloroplast count (Supp. Fig. 2.2). When normalizing to either

chlorophyll or chloroplast count, no changes in mean ion concentration were statistically

significant. However, we did observe some promising trends when normalizing element

concentrations to total chlorophyll. As in the leaf ionome data, opt3-2 had higher levels of Fe than

both the WT and fro7 mutant. Unlike in leaf tissue, Mn levels were equivalent between the

genotypes. In opposition to our expectations, chloroplasts from fro7 did not exhibit reduced Fe

levels as was previously published (Jeong et al., 2008). Yet, fro7 exhibited lower levels of K and

Ca, in opposition with what we observed in leaf tissue, where fro7 accumulated more K and Ca.

These trends merit further investigation by repetition of experiments.

Preliminary Data: TXRF measurement and analysis reveals kea1kea2 chloroplasts accumulate

higher concentrations of several elements.

As for WT, fro7, and opt3-2 chloroplasts, we normalized element concentrations in kea1kea2 to

both total chlorophyll (Supp. Fig. 2.3) and chloroplast count (Fig. 2.6). In this case, our preferred

method of normalization was to chloroplast count, as kea1kea2 mutants are chlorophyll deficient.

Remarkably, chloroplasts from both independent kea1kea2 lines accumulated not only more K

than WT chloroplasts isolated simultaneously (black points for WT chloroplasts isolated alongside

kea1-1kea2-1, and red points for chloroplasts isolated alongside kea1-2kea2-2) but also more Ca,

Mn, Fe, Ni, and Cu. Previous data collected with nitric acid digested chloroplasts (Supp. Fig. 2.4)

from WT and both kea1kea2 lines also shows a similar trend towards over-accumulation of all

elements in kea1kea2, although absolute concentrations are much different. While further

experiments are required to see if these trends are repeatable, this preliminary data indicates that

loss of KEA1/2 transporters dramatically alters the whole chloroplast ionome.

38

Fig. 2.6: Elemental analysis of plastids from WT and kea1kea2 lines using TXRF. Concentration of A potassium (K), B calcium (Ca), C manganese (Mn), D iron (Fe), E nickel (Ni), F copper (Cu), and G zinc (Zn) normalized to chloroplast count (ng*106 chloroplasts-1, n = 1-2). Points with corresponding color represent replicates isolated on the same day.

39

DISCUSSION

Using a modified chloroplast isolation protocol and establishing procedures for preparation and

measurement of isolated chloroplasts with TXRF analysis, we were able to quantify many

biologically relevant elements simultaneously in isolated chloroplasts. While this method requires

further improvement with regards to precision and reproducibility, the concentrations of many

elements, particularly transition metals Mn, Fe Cu, and Zn, were very similar to the literature.

Other elements, including Ca and K, have never been measured in A. thaliana chloroplasts. Further

experiments and verification of concentrations with well-established alternative methods such as

atomic absorption spectroscopy (AAS), Inductively Coupled Plasma Atomic Emission

Spectroscopy (ICP-AES) and/or Inductively Coupled Plasma Mass Spectrometry (ICP-MS) are

necessary to validate our results.

However, with our method as it is, we were able to quantify some interesting trends in

elemental accumulation in isolated chloroplasts from the fro7, opt3-2, and kea1kea2 mutants in

comparison to the wildtype. Notably, our method revealed that opt3-2, which has been previously

published to over-accumulate iron in leaf tissue, also accumulates high levels of iron in the

chloroplast. The OPT3 transporter localizes to the plasma membrane where it participates in iron-

loading into the phloem (Zhai et al., 2014). This protein is also linked to shoot-to-root Fe sensing,

which results in knockdown lines exhibiting constitutively high expression of Fe uptake proteins

in the roots, and subsequently accumulation of high levels of Fe in leaf tissue (Stacey et al., 2008;

Zhai et al., 2014). Considering that a large portion of leaf Fe is stored in the chloroplast (Kroh and

Pilon, 2020), it is reasonable that opt3-2 chloroplasts had roughly 3-fold more Fe than WT

chloroplasts. While Fe is an essential cofactor in almost every complex of the plastid Electron

40

Transport Chain (ETC), free Fe can interact with hydrogen peroxide (H2O2) to produce dangerous

hydroxyl free radicals (•OH) (Kroh and Pilon, 2020; Schmidt et al., 2020). These hydroxyl radicals

can then damage numerous biological molecules including components of the ETC (Pilon et al.,

2011). Fe-mediated •OH production can even trigger cell death, although so far this has only been

shown in root cells (Kazan and Kalaipandian, 2019). A large amount of H2O2 is produced in the

chloroplast from superoxide generated by the plastid ETC (Smirnoff and Arnaud, 2019). Thus, Fe

excess in the chloroplast and subsequent reaction with H2O2 may account for the dwarf phenotype

and necrotic lesions exhibited by the opt3-2 mutant.

The chloroplasts from the fro7 mutant lack a Fe(III) chelate reductase thought to be

required for Fe3+ reduction and subsequent uptake into the plastid. When analyzed with our

method, fro7 chloroplasts did not show the expected decrease in plastid Fe concentration (Jeong

et al., 2008). However, the published decrease in fro7 chloroplast iron content compared to the

WT was only 33%, thus our method may be too imprecise at present to resolve such a small change.

Furthermore, it has been shown that increasing overall Fe treatment can ameliorate the fro7

phenotype. Therefore, our plants may not show an iron deficiency in leaves or isolated plastids

due to there being sufficient Fe in the soil to compensate for the loss of FRO7. However, we did

observe some interesting trends towards decreased K and Ca concentration in fro7 chloroplasts.

This was in opposition to our measurements of fro7 leaf tissue, which had higher levels of these

elements compared to the WT. Recent publications have linked Fe and K homeostasis, suggesting

that K promotes Fe translocation from roots to shoots, and also exacerbates Fe toxicity (Çelık et

al., 2010; Ye et al., 2019). This phenomenon may be tied to altered leaf K concentrations in both

opt3-2 and fro7. However, the direct connection between subcellular K and Fe homeostasis is far

from clear and merits more investigation. As for leaf and plastidial Ca and Fe concentrations, there

41

is no clear link in the literature. Further investigation of the interaction between Fe and Ca

homeostasis is needed.

After testing proof-of-concept with WT, fro7, and opt3-2, we accomplished some preliminary

characterization of the chloroplast ionome of plastid K+/H+ antiporter mutant kea1kea2. We

hypothesized that this mutant would accumulate more K in the chloroplast than the WT, and our

data supported this hypothesis so far. Isolated chloroplasts from kea1kea2 had 2-fold higher K than

the WT, yet additionally had higher levels of Ca, Fe, Ni, and Cu. This suggests that alteration of

K+ transport influences overall chloroplast ion homeostasis.

CONCLUSIONS

We used our kea1kea2 chloroplast ionome results to begin to build a model of how loss of K+/H+

transport across the plastid envelope influences photosynthesis and chloroplast development (Fig.

2.7). In a WT leaf cell (Fig. 2.7A), the proton gradient across the chloroplast envelope drives H+-

coupled K+ efflux from the stroma to the cytosol via KEA1 and KEA2 antiporters. This maintains

moderate levels of K in the plastid and promotes overall ion homeostasis. WT chloroplasts develop

normally and do not exhibit any defects. In contrast, chloroplasts in the kea1kea2 leaf cell (Fig.

2.7B) lack KEA1 and KEA2 proteins due to large T-DNA insertions which disrupt transcription

of both genes. Thus, kea1kea2 chloroplasts accumulate high levels of K in the stroma, which may

perturb the homeostasis of other elements. The downstream effects of excess K and other elements

will be investigated in the next chapters.

42

METHODS

Plant growth conditions. Plants were grown as follows; seeds were sprinkled onto pots of damp

potting mix, then vernalized for 48 hours in the dark at 4° C. Seeds on soil were then placed in a

Percival Growth Chamber with 150 µmol photons m-2s-1 of light, 16 h: 8 h light: dark cycle.

Fig. 2.7: Model of how loss of K+/H+ exchangers KEA1/2 influences leaf plastid ionome. A In a WT cell exposed to light with both KEA1 and KEA2, potassium accumulates in the plastid through an unknown mechanism. Simultaneously, light-driven proton pumping from the stroma to the thylakoid lumen increases stromal pH, making it basic (pH ~8) compared to the cytosol (~pH 7). This pH gradient causes KEA1 and KEA2 to antiport protons into the stroma in exchange for K+ ions, thus acting as a valve for K+. B In an illuminated kea1kea2 leaf cell, the TDNA insertions in KEA1 and KEA2 result in complete loss of KEA1/2 antiporters. As a consequence, K+ ions which enter the plastid have limited mechanisms for exiting the plastid. This results in excessive accumulation of K+ in the plastid, which presumably is causal for the chlorotic phenotype and developmental delay in kea1kea2 plastids.

43

Growth temperatures were 22° C in the light and 18° C in the dark. Plants were grown for 21-28

days until harvest.

Plant mutant isolation and information. WT (Col-0, CS70000) seeds were obtained from the

ABRC. Homozygous kea1-1kea2-1 and kea1-2kea2-2 were obtained from a previous study (Kunz

et al., 2014). Fro7 and opt3-2 were generously provided by Dr. David Mendoza-Cózatl (University

of Missouri).

Accession Numbers for this study. KEA1 (AT1G01790), KEA2 (AT4G00630), FRO7

(AT5G49740), and OTP3 (AT4G16370).

Pulse-Amplitude-Modulation (PAM) fluorescence spectroscopy. Chlorophyll fluorescence

measurements were taken as previously described (Kunz et al., 2009). Plants were dark adapted

for 20 minutes prior to imaging. Imaging was carried out on an Imaging PAM M-series chlorophyll

fluorometer (Walz) using a built-in induction protocol for chlorophyll fluorescence kinetics with

a photosynthetically active radiation (PAR) intensity of 185 photons m-2s-1.

Chlorophyll quantification. Chlorophyll was quantified as described in (Porra et al., 1989). In

brief, leaf tissue was ground to a fine powder in liquid nitrogen. Chlorophyll from 10 mg of leaf

powder was extracted by vortexing vigorously in 5 mL of ice-cold 80% acetone. Samples were

incubated in the dark for 2 hours in a sonic bath filled with ice water, then were centrifuged at 4°

C and maximum speed for 10 minutes. For isolated chloroplasts, 10 µL were diluted in 1990 µL

of ice-cold 80% acetone, vortexed vigorously, and centrifuged at 4° C and maximum speed in a

benchtop centrifuge. Optical density (OD) of the supernatant was then measured in a

spectrophotometer at 646, 663, and 750 nm. Chlorophyll A and B concentrations were calculated

44

using the OD750 as the baseline for scatter. For leaf tissue samples, chlorophyll levels were

normalized to fresh weight.

Chloroplast isolation and evaluation. Chloroplasts were isolated using a modified version of the

protocol described in (Aronsson and Jarvis, 2002). 5 g of leaves were harvested from 3-4 week old

plants. Leaves were then washed 3 times in distilled water and placed at 4°C for 10 minutes to

inactivate native proteases. Then, 2.5 g of leaves were ground with 25 mL of isolation buffer (0.3

M sorbitol, 5 mM MgCl2, 5 mM EDTA, 20 mM HEPES, 10 mM NaCO3, 45 mM ascorbic acid,

pH 8.0 with NaOH) in a chilled warring blender with six one-second pules on high speed. Lysate

was then filtered through a layer of fine cheesecloth. The grinding process was repeated for the

other 2.5g of leaves. Homogenate was then centrifuged at 4° C and 15,000g for 4 minutes.

Supernatant was discarded, and pellet was gently resuspended in 1 ml of isolation buffer using a

fine paintbrush, and layered over a Percoll step gradient (30%, 82% Percoll in 0.3 M sorbitol 5

mM MgCl2, 5 mM EDTA, 20 mM HEPES-NaOH pH 8.0) and centrifuged at 2000g for 12 minutes

with no brake. The lower band containing intact chloroplasts was carefully removed, then mixed

in 3 volumes of wash buffer (50 mM HEPES, 0.3 M sorbitol, pH 8.0 with NaOH) and centrifuged

for 4 minutes at 1500g and 4 ºC. Supernatant was discarded, and wash step was repeated once

more. Pelleted chloroplasts were then resuspended with a fine paint brush in a small volume of

wash buffer. For chlorophyll quantification was carried out as described above. For counting and

evaluation of intactness, chloroplasts were diluted 10-fold with wash buffer, applied to a Hauser

cell-counting chamber, and viewed at 20x magnification on a light microscope under phase-

contrast settings.

45

Sample preparation for TXRF. Chloroplasts were mixed with Gallium (Ga) and Scandium (Sc)

internal standards, Polyvinyl Alcohol (PVA) and water to make a final concentration of 100-400

µg/mL chlorophyll, 1 part per million (ppm) Ga, 1 ppm Sc, and 0.2% PVA. As a control, an

equivalent volume of wash buffer was prepared as described. For leaf samples, ~10 mg of dry,

powdered material was digested in 500 µL of HNO3 at 90° C for 1 hour. Digest was diluted 10-

fold with deionized water and internal standards to make a final concentration of 1 ppm Ga and 50

ppm Sc. Samples were then spotted on a silicone-coated quartz carrier and dried under vacuum or

on a hot plate.

TXRF measurement and analysis. Once dry, carriers were placed into a Bruker T-star S4. Lighter

elements (P to V) were measured using the tungsten L-line x-ray beam with Sc as the internal

standard. Heavier elements (Mn to Zn) were measured using the molybdenum K-line X-ray beam

with Ga as the internal standard. With both excitations, fluorescence was counted for 600 and 1000

seconds for leaf tissue and chloroplasts, respectively. Peak deconvolution was done by Bruker

Tspirit software. Element concentrations measured in chloroplast or tissue-free blanks were

subtracted from concentrations measured in biological samples. Chloroplast element

concentrations were then normalized to chloroplast count or chlorophyll content. Leaf tissue

element concentrations were normalized to dry weight (DW).

Statistical Analyses. All statistical tests were done in Graphpad Prism™ version 8. Bioreplicate

values were determined to be normally distributed or not based on a Shapiro-Wilk test. Means for

data points that were normally distributed were tested for statistical significance using one-way

ANOVA with Geisser-Greenhouse correction and Holm-Sidaks multiple comparisons test. Means

for data points that were not normally distributed were tested for statistical significance using a

46

Friedman test and Dunn’s multiple comparisons test. Means whose p-value was less than 0.05

were considered statistically significantly different.

47

SUPPLEMENTAL TABLES

Supp. Table 2.1: Element Analysis of NIST reference material Spinach 1570a Tomato 1573a Apple 1515

Exp. conc.

Mean Conc.

Std. Dev. % R

Exp. conc.

Mean Conc.

Std. Dev. % R

Exp. conc.

Mean Conc.

Std. Dev. % R

Na 18.210

166.50

157.69

>200 0.136 160.88

207.03

>200 0.024 160.56

8.531 >200

Mg 9.000 5.151 0.109 57 12.000

6.230 0.506 52 2.710 1.981 0.941 73

Al 0.31 3.01 1.05 >200 0.60 3.53 0.94 >200 0.28 1.95 0.07 >200

P 5.19 6.62 0.52 128 2.16 3.27 0.06 152 1.59 1.92 0.23 120

S 5.00 5.55 0.71 111 9.60 11.60 0.43 121 1.80 2.17 0.39 121

Cl N/A 1.65 0.23 N/A 6.60 1.26 0.09 19 0.58 0.08 0.01 15

K 29.00 27.46 1.40 95 26.76 27.14 0.83 101 16.08 14.81 2.35 92

Ca 15.26 19.45 9.22 127 50.45 57.73 0.95 114 15.25 16.91 2.34 111

Mn 0.076 0.0791

0.0061

104 0.2463

0.2489

0.0009

101 0.0541

0.0488

0.0079

90

Fe N/A 0.2919

0.0750

N/A 0.3675

0.3370

0.0440

92 0.0827

0.0681

0.0121

82

Ni 0.0021

0.0031

0.0021

143 0.0016

0.0019

0.0007

119 0.0009

0.0010

0.0007

103

Cu 0.0122

0.0127

0.0022

104 0.0047

0.0041

0.0023

88 0.0057

0.0085

0.0030

149

Zn 0.082 0.088 0.209 107 0.031 0.301 0.181 >200 0.012 0.321 0.251 >200

Rb N/A 0.0127

0.0010

N/A 0.0148

0.0157

0.0014

106 0.0102

0.0092

0.0015

90

Sr 0.0555

0.0540

0.0057

97 0.0850

0.0831

0.0007

98 0.0250

0.0226

0.0032

90 Expected element concentration, and mean element concentration measured using Total-Reflectance X-RAY Fluorescence (TXRF) of National Institutes of Standards and Technology (NIST) reference materials in mg per g dry weight (mg * g-1 DW). Percent recovery (% R) was calculated as 100*(measured concentration)/ (expected concentration). Analysis was conducted on 3 separate technical replicates of each material which were independently weighed, digested in nitric acid, diluted, and measured.

Supp. Table 2.2: TXRF measurements of standards spiked into isolated chloroplasts Water Chloroplast Buffer Chloroplasts

Std.

Exp. conc. (ppm)

Mean Conc. (ppm)

Std. Dev.

(ppm) % R

Mean Conc. (ppm)

Std. Dev.

(ppm) % R

Mean Conc. (ppm)

Std. Dev.

(ppm) % R

V 20 21.233 0.142 106 28.137 1.038 141 21.187 1.269 106 2 2.140 0.020 107 2.046 0.029 102 1.894 0.033 95

0.2 0.218 0.007 109 0.186 0.002 93 0.177 0.003 88

Rb 20 20.617 0.802 103 22.387 0.468 112 21.847 1.841 109 2 2.254 0.014 113 2.111 0.018 106 2.013 0.048 101

0.2 0.230 0.010 115 0.212 0.001 106 0.209 0.008 105 Expected element concentration, and mean element concentration measured using Total-Reflectance X-RAY Fluorescence (TXRF) of elements spiked into samples of water, chloroplast buffer, and isolated chloroplasts prepared with PVA and internal standards as described in the methods section. Chloroplast samples had a final chlorophyll concentration of 400 µg*mL-1). Measurements of mean element concentration and standard deviation of the mean (Std. Dev) are in parts per million (g*L-1). Percent recovery (% R) was calculated as above. Analysis was conducted on 2-3 separate technical replicates of each material which were independently pipetted and measured.

48

SUPPLEMENTAL FIGURES

Supp. Fig. 2.1: A sample of chloroplasts digested in HNO3. Plastids were then spotted on a TXRF quartz carrier and dried. Note puffy appearance and thickness of sample.

49

Supp. Fig. 2.2: Analysis of plastids from WT, fro7, and opt3-2 norm. to chlorophyll. Mean concentration of A potassium (K), B calcium (Ca), C manganese (Mn), D iron (Fe), E nickel (Ni), F copper (Cu), and G zinc (Zn) normalized to normalized to chloroplast count (ng*10-6 chloroplasts, +/- SD, n = 3). No statistically significant differences between genotypes were detected (p < 0.05). Points with corresponding color represent replicates isolated on the same day.

50

Supp. Fig. 2.3: Analysis of plastids from WT and kea1kea2 norm. to chlorophyll. Concentration of A potassium (K), B calcium (Ca), C manganese (Mn), D iron (Fe), E nickel (Ni), F copper (Cu), and G zinc (Zn) normalized to total chlorophyll (ng*µg Chl-1, n = 1-2). Points with corresponding color represent replicates isolated on the same day.

51

Supp. Fig. 2.4: Elemental analysis of HNO3-digested plastids. Concentrations normalized to chloroplast count (ng*10-6 chloroplasts, n = 4).

Potass

ium

Calci

umIro

n

Copper

0

50

100

150

200300400

Abundant Elementsn

g*1

0-6 c

hlo

rop

las

ts

kea1-2kea2-2

WT

kea1-1kea2-1

Man

ganes

e

Nicke

lZin

c

0

2

4

6

101520

Rare Elements

ng

*10-6

ch

loro

pla

sts

kea1-2kea2-2

WT

kea1-1kea2-1

52

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Kunz, H.H., Gierth, M., Herdean, A., Satoh-Cruz, M., Kramer, D.M., Spetea, C., and Schroeder, J.I. (2014). Plastidial transporters KEA1, -2, and -3 are essential for chloroplast osmoregulation, integrity, and pH regulation in Arabidopsis. Proc Natl Acad Sci U S A. 111, 7480-7485.

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Stacey, M.G., Patel, A., McClain, W.E., Mathieu, M., Remley, M., Rogers, E.E., Gassmann, W., Blevins, D.G., and Stacey, G. (2008). The Arabidopsis AtOPT3 protein functions in metal homeostasis and movement of iron to developing seeds. Plant Physiol. 146, 589-601.

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Provide Insights on the Effects of pH and Ion Homeostasis. Plant Physiol. 177, 875-895. Terry, N., and Low, G. (1982). Leaf chlorophyll content and its relation to the intracellular

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56

CHAPTER 3: PERTURBATION OF CHLOROPLAST ION TRANSPORT INFLUENCES NUCLEAR AND PLASTID GENE EXPRESSION

Figures and text in this chapter were submitted for publication in The Plant Cell with coauthors Rouhollah Barahimipour, Nikolay Manavski, Ricarda Höhner, Serena Schwenkert, Bettina Bölter, Takehito Inaba, Jörg Meurer, Reimo Zoschke and Hans-Henning Kunz.

ABSTRACT

The K+/H+ exchangers KEA1 and KEA2 of the inner plastid envelope are critical for

chloroplast development, ion homeostasis and proper photosynthesis. However, the exact

mechanism by which loss of KEA transporters affects organelle biogenesis remains elusive.

Chloroplast development requires intricate coordination between the nuclear and plastid

genome (plastome). Many mutants compromised in plastome gene expression display a

virescent phenotype, i.e., delayed greening. The phenotypic appearance of kea1kea2 double

mutants fulfills the same criteria, yet a link to plastid gene expression has not been explored.

Here, we used a systems-biology approach to uncover evidence in nuclear transcriptomics

that plastid RNA metabolism is altered due to loss of KEA1/2. A closer look at plastome

RNAs confirmed that loss of KEA1/2 results in maturation defects of the plastid ribosomal

RNAs. Mutant plants also exhibit hampered protein synthesis and lower steady-state levels

of plastome-encoded proteins. Hence, the loss of K+/H+ antiporters does indeed impair plastid

gene expression on the post-transcriptional, and translational level, which is likely the reason

for delayed chloroplast development. Additionally, neither expression of nuclear-encoded

plastid RNA-binding proteins nor plastid protein import in kea1kea2 is disrupted. These

results provide evidence for a new hypothesis; Disturbed stromal ion homeostasis prevents

the activity or binding of plastid RNA-binding proteins to their RNA targets. This effect

and/or the negative impact of altered stromal ion levels on ribosomal assembly potentially

57

culminate in the reduced plastid translation rates we quantified and cause delayed

chloroplast development.

INTRODUCTION

An overview of potassium (K) transport proteins.

Plants have evolved numerous genes involved exclusively in the uptake and transport of charged

molecules such as ions across cell and organelle membranes. In model plant species Arabidopsis

thaliana, 2411 of the 27655 annotated genes in the genome are associated with transport-related

functions (Tair10 GO ontology). Mineral nutrients are typically not present in high concentrations

in the substrate, so many plant transporters are involved in the uptake and bulk transport of

nutrients for use in biomolecules. Transport proteins are also essential to maintain the

characteristic ion and pH compositions compartmentalized in different membrane-bound

organelles, i.e., subcellular ion homeostasis (Sze and Chanroj, 2018). Due to the high abundance

of potassium (K) and its unique importance for moderating pH, plants have evolved a slew of

transport mechanisms for this ion with different properties suited for high and low affinity uptake

from media, long distance vascular transport, import into cells, and lastly import into subcellular

compartments (Ragel et al., 2019). In A. thaliana, several dozen genes spread across 4 families are

responsible for uptake and maintenance of cellular and organellar K gradients (Fig. 3.1) (Hedrich,

2012; Sharma et al., 2013; Ragel et al., 2019). These include two families of channels, one family

of symporters, and one family of proton-coupled antiporters, known as the Cation Proton

Antiporter (CPA) family (Mäser et al., 2001; Ragel et al., 2019). The CPA family is the largest of

58

Fig. 3.1: An overview of K transport protein in A. thaliana. The rough phylogenetic relationship, function, and members of the 5 families of A. thaliana K transport proteins. Plasma membrane = PM, vacuolar membrane = VM, Endomembrane = EM, plastid (chloroplast) membrane = CM. Images of KEA proteins from Tsujii et al. 2019.

59

the four families and plays a critical role in subcellular ion and pH homeostasis (Pittman, 2012;

Sze and Chanroj, 2018). Thus, these carriers are potentially very important for the function of

bioenergetic organelles such as the mitochondria and the chloroplast, which rely on carefully

moderated proton-motive force (PMF) across membranes to generate biomolecules. However,

within the CPA superfamily, only 3 K+/H+ carriers have been definitively localized to a

bioenergetic organelle; three of the six K Efflux Antiporter (KEA) proteins are targeted to the

plastid (Aranda-Sicilia et al., 2012; Kunz et al., 2014). Other subfamilies including NHX, SOS,

and CHX type transporters are either Na-specific, or do not localize to plastid membranes (Sze

and Chanroj, 2018). Unlike other CPA protein subfamilies, KEA transporters are highly

homologous to cyanobacterial transporter KefC, with no close animal, fungal, or protozoan

homologues (Chanroj et al., 2012). This indicates the KEA family may have evolved specifically

within the green chloroplast from a cyanobacterial endosymbiont. All KEA transporters contain

10 transmembrane helices and show specificity for transporting K+ rather than other cations such

as Na+ (Tsujii et al., 2019; Aranda Sicilia et al., 2021). However, KEA4, 5, and 6 localize to the

endomembrane system or prevacuole, and are most highly expressed in roots, anthers, and stem

vasculature (Klepikova et al., 2016; Zhu et al., 2018). KEA1, 2, and 3 have a similar

transmembrane sequence as KEA4-6, yet additionally contain an N-terminal chloroplast transit

peptide, and a C-terminal KTN domain related to transporter regulation (Bölter et al., 2019; Tsujii

et al., 2019). Within the chloroplast, KEA1 and KEA2 localize to the inner envelope, while KEA3

localizes to the chloroplast thylakoid membrane (Kunz et al., 2014). All three genes are most

highly expressed in young leaf tissue (Klepikova et al., 2016). As was described in the first chapter,

KEA3 has a well-characterized role in maintaining PMF partitioning to regulate non-

photochemical quenching (Armbruster et al., 2016; Wang et al., 2017). A. thaliana loss-of-function

60

mutants for KEA3 have no obvious phenotype unless grown under fluctuating light, a condition

where rapid NPQ induction and relaxation is required for optimal growth. In contrast, a

simultaneous loss of both functionally redundant KEA1/2 antiporters results in a stunted plant with

elevated leaf K content (Höhner et al., 2016b; Höhner et al., 2019). The direct impacts of these

transporters on bioenergetic processes within the chloroplast such as PMF are well documented

(Kunz et al., 2014). Additionally, a recent study showed that KEA1 and KEA2 are also important

for chloroplast biogenesis as the double mutants display chlorotic young leaves with delayed

greening and immature chloroplasts (Aranda-Sicilia et al., 2016). Studies with Oryza sativa

homologue AM1 reveal a similar role for the protein family in monocot lineages (Sheng et al.,

2014). However, the mechanistic role of ion transport via KEA transporters in plastid development

remains an outstanding scientific question (Sze and Chanroj, 2018). Thus, the objective of this

chapter is to elucidate the connection between KEA-mediated ion/proton transport and

chloroplast development.

Chloroplast development and the complicated process of plastid gene expression (PGE).

Chloroplasts develop during the plant life cycle from a photosynthetically inactive proplastid

through coordinated expression of specific genes (Pogson and Albrecht, 2011; Hernandez-Verdeja

and Strand, 2018). This coordination is complex because plastids have retained a small genome of

~120 genes known as the plastome, which is critical for chloroplast function (de Vries and

Archibald, 2018). The plastome encodes many components of the electron transport chain,

including the photosystem reaction centers (Allen et al., 2011). The majority of chloroplast genes

were transferred to the nuclear genome, including the light harvesting machinery and chlorophyll

biosynthesis enzymes, a group collectively known as the Photosynthesis-Associated Nuclear-

Encoded Genes (PhANGs), reviewed in Berry et al., 2013. Notably, many photosynthetic protein

61

complexes including the photosystems, the cytochrome b6f complex, ATP synthase, and Rubisco

contain subunits from both genomes (Allen et al., 2011).

The division of genes encoding the chloroplast proteome between the nuclear genome and

the plastome necessitates tight regulation and communication between the nucleus and the

chloroplast to balance protein stoichiometry (Woodson and Chory, 2008). Nucleus to plastid

communication to moderate plastome gene expression (PGE) is known as anterograde signaling.

Anterograde signaling impacts plastid gene expression at the transcriptional, posttranscriptional,

translational, and post-translational level via nuclear-encoded proteins. In particular, chloroplast

RNA-binding proteins (cRBPs) play a key role in posttranscriptional and translational regulation

(Barkan and Small, 2014; Zoschke and Bock, 2018). After translation in the cytosol, chloroplast

RNA-binding proteins are imported into the plastid where they assist in stabilizing, end processing,

splicing, and editing of mRNAs and rRNAs (Schmitz-Linneweber and Small, 2008; Hammani et

al., 2014). A slew of mutants defective in anterograde control of PGE including pentatricopeptide

repeat proteins (PPRs ,e.g. ys1, opt70, rap1, Zhou et al., 2009; Chateigner-Boutin et al., 2011;

Kleinknecht et al., 2014; Zoschke et al., 2016), ribosomal proteins (e.g. rps5, Zhang et al., 2016),

chloroplast translation initiation and elongation factors (e.g. fug1, svr11; Miura et al., 2007; Liu

et al., 2019), and proteases (e.g. clpt1clpt2 , Kim et al., 201) have been isolated and characterized

over the last decade. Intriguingly, many of these mutants display a distinctive “virescent” (delayed-

greening) phenotype (Zhou et al., 2009). In virescent mutants, young leaves remain pale with poor

rates of photosynthesis, whereas older leaves have close to wild-type (WT) levels of chlorophyll

and photosynthesis. A selection of these mutants growing alongside kea1kea2 is shown in Fig. 3.2.

The afore-mentioned pale young leaf phenotype in kea1kea2 double mutants appears to phenocopy

many “virescent” A. thaliana mutants with defects in plastid gene expression (Fig. 3.2). However,

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it has not been investigated if the virescent phenotype exhibited by kea1kea2 results from changes

in PGE. Thus, our operating hypothesis is that loss of K+/H+ homeostasis due to the absence

of KEA1/2 disrupts PGE, and thereby impairs plastid development and photosynthesis.

Intriguingly, the virescent phenotype in kea1kea2 mutants can be suppressed by exposing plants

to NaCl stress (Kunz et al., 2014). This behavior suggests kea1kea2 may serve as a useful model

to gain insights into the relationship between chloroplast ion homeostasis and PGE. Furthermore,

investigation of PGE may also provide a mechanistic explanation for the delayed chloroplast

biogenesis documented in kea1kea2 loss-of-function lines (Aranda-Sicilia et al., 2016), and answer

one of the outstanding questions in the field (Sze and Chanroj, 2018). Using transcriptomics

combined with the NaCl recovery phenomenon in kea1kea2, we set out to gain insights into

the molecular basis of the kea1kea2 phenotype and its potential connection to PGE.

Fig. 3.2: kea1kea2 double mutants phenocopy many PGE mutants. Top panels are RGB photos showing delayed greening in kea1kea2 and PGE mutants. Bottom row of panels shows reduced Fv/Fm

in young leaves of kea1kea2 and assorted PGE mutants.

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RESULTS

Loss-of-function lines for kea1kea2 over-accumulate potassium, which can be prevented by

exogenous sodium treatment.

To begin, we investigated how the leaf ionome changes in response to exogenous NaCl

application, which has been previously shown to suppress the photosynthetic phenotype in

kea1kea2(Kunz et al., 2014). It was published previously that loss of KEA1/2 function in plants

growing on regular media results in increased K concentrations in leaf tissue (Table 3.1, Höhner

et al. 2016). However, data regarding how exogenous sodium treatment influences K levels or

other ions has not been published. Thus, we re-analyzed some TX-RF ionome data generated by

Dr. Ricarda Höhner, which included WT plants and kea1kea2 lines grown with or without

exogenous 75mM NaCl treatment. Under control conditions, both independent kea1kea2 lines

accumulated significantly more potassium (K) and phosphorus (P), yet significantly less

magnesium (Mg, Table 3.1 bolded values). There were also changes in the accumulation of other

elements (e.g. Mangenese, Mn) but these changes were either only statistically significant in one

kea1kea2 line, or weren’t statistically significant in either line. When subjected to NaCl treatment,

the WT and both kea1kea2 lines accumulated lower levels of K in leaves compared to the control

condition (Table 3.1). Notably, K concentrations in kea1kea2 were no longer significantly

increased compared to WT under NaCl treatment. Application of NaCl had no influence on the

accumulation of Mg in the WT, but increased Mg levels in kea1kea2 to WT levels. The only

element that did not reliably return to WT-levels under NaCl treatment was P, which was lower in

kea1-1kea2-1, but remained high in kea1-2kea2-2. Naturally, the accumulation of several other

elements (e.g., Na, Cl) was increased or otherwise altered under NaCl treatment, but these changed

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uniformly across all genotypes and thus were likely not the related to the kea1kea2 phenotype. In

general, the recovery of the kea1kea2 leaf ionome under NaCl treatment mirrors the photosynthetic

recovery shown in the literature (Kunz et al., 2014). This opens the question as to if the altered

accumulation of one or more of the elements is causal for the photosynthetic phenotype. The two

elements which showed consistent recovery to WT levels under NaCl treatment, K and Mg, are

prime candidates. Little has been published about K over-accumulation in the literature, as this

type of stress is rare in nature or in agriculture. However, Mg deficiency has been shown to reduce

photosynthetic efficiency and result in chlorosis (Hermans and Verbruggen, 2005; Farhat et al.,

2016). Intriguingly, the concentration of the two elements is linked, as excess K can induce Mg

deficiency by out competition for transport (Marschner, 1995). To investigate further, we grew

all three genotypes on media with added KCl to see if additional K exacerbates the kea1kea2

phenotype (Fig. 3.3), and with additional MgCl2 to see if added Mg rescues the kea1kea2

Table 3.1: Leaf-level concentrations of elements (µg*mg DW -1)

WT kea1-1kea2-1 kea1-2kea2-2

Control NaCl Control NaCl Control NaCl

Mean Std Err. Mean

Std Err. Mean

Std Err. Mean

Std Err. Mean

Std Err. Mean

Std Err.

Na 0.54 0.11 14.56 2.23 1.23 0.33 15.89 3.07 1.30 0.45 17.74 1.84

Mg 5.78 0.76 5.34 0.20 3.10 0.34 5.31 0.18 3.69 0.24 4.77 0.62

P 8.10 0.53 7.13 0.35 11.54 0.69 11.11 2.10 11.96 0.40 7.82 0.21

S 4.37 0.58 7.06 0.32 5.15 0.31 8.20 0.87 6.30 0.90 6.64 0.32

Cl 1.22 0.08 11.72 1.40 1.15 0.11 10.66 0.89 1.40 0.29 11.89 1.28

K 40.59 2.15 29.37 1.39 51.68 0.99 31.91 1.95 53.11 1.80 29.88 0.76

Ca 36.07 3.09 32.27 1.52 41.83 8.72 28.85 1.61 34.36 4.29 34.61 6.12

Mn 0.08 0.00 0.08 0.00 0.12 0.02 0.09 0.01 0.10 0.01 0.11 0.01

Fe 0.14 0.02 0.12 0.01 0.22 0.05 0.09 0.01 0.18 0.05 0.15 0.03

Cu 0.01 0.00 0.01 0.00 0.02 0.00 0.01 0.00 0.06 0.04 0.01 0.00

Zn 0.73 0.13 0.50 0.10 1.02 0.29 0.51 0.07 1.05 0.45 0.39 0.07

Rb 0.01 0.00 0.01 0.00 0.01 0.00 0.01 0.00 0.01 0.00 0.01 0.00

Sr 0.06 0.01 0.05 0.00 0.07 0.01 0.05 0.01 0.06 0.01 0.08 0.02

Pb 0.15 0.04 0.11 0.03 0.18 0.03 0.06 0.02 0.16 0.04 0.09 0.02 Mean leaf concentration of assorted elements normalized to dry weight (µg*mg DW -1) and standard error of the mean (Std. Err, n =6-7). Bolded values indicate statistically significantly different means from correspondingly treated WT sample based on Dunnett's or Dunn's multiple comparison test (p < 0.05).

65

phenotype (Supp. Fig. 3.1). We also grew these lines on media with additional NaCl to ensure we

could reproduce the effect observed previously.

Sodium treatment ameliorates delayed greening, and improves photosynthetic yields in kea1kea2,

while potassium treatment exacerbates it.

Next, we evaluated the phenotype of the kea1kea2 mutant on regular ½ MS media, and media

supplemented with KCl, MgCl2, or NaCl. With the naked eye, we observed that loss of KEA1/2 in

plants grown on regular media results in decreased biomass and chlorosis (Fig 3.4A control). The

addition of 75 mM KCl decreased the biomass of kea1kea2 lines and the WT (Fig 3.4A KCl). As

shown previously, the addition of 75 mM NaCl to growth media recovered chlorophyll in the

kea1kea2 mutant but did not increase biomass (Fig. 3.3A ‘NaCl’, Kunz et al., 2014). Evaluation

of photosynthetic parameters using Pulse Amplitude Modulated (PAM) fluorescence (see

Appendix A for theoretical overview of this method) replicated previous results showing the

KEA1/2 loss-of-function line exhibits low maximum quantum yield of photosystem II (Fv/Fm, Fig.

3.3 A-B ‘control’, Kunz et al 2014). The false-color image of Fv/Fm (Fig. 3.3A, bottom panel)

shows the typical virescent phenotype of kea1kea2, where young leaves exhibit the greatest

impairment of photosynthesis. Loss of KEA1/2 under control conditions also results in altered

partitioning of captured light energy. In kea1kea2, significantly more harvested light energy is

routed through regulated nonphotochemical quenching (ΦNPQ) and non-regulated non-

photochemical quenching (ΦNO) than through photosystem II photochemistry (ΦII) compared to

the WT (Fig. 3.3C-E). KCl treatment further decreased Fv/Fm in kea1kea2 compared to the KCl-

treated WT (Fig 3.4A-B ‘KCl’). In kea1kea2 under excess K stress, energy partitioning to ΦNO

increased even more with concurrent decreases in ΦII. In contrast, NaCl treatment increased Fv/Fm

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Fig. 3.3: KCl treatment exacerbates the kea1kea2 photosynthetic phenotype, while NaCl rescues phenotype. (A) Phenotype of WT and kea1kea2 lines grown without (control), with 75 mM KCl, or with 75 mM NaCl. Lower panels display false-color image of maximum quantum yield of PSII (Fv/Fm). (B-E) Bar graph of mean photosynthetic parameter in WT and kea1kea2 under control (white bars), KCl treatment (light grey bars), or NaCl (dark grey bars, ± SEM, n = 40-44). Parameters include (B) theoretical maximum yield of PSII, Fv/Fm (C) flux through PSII, ΦII (D) flux through regulated Non-photochemical quenching, ΦNPQ, and (E) flux through non-regulated, non-photochemical quenching, ΦNO (Kramer et al., 2004; Klughammer and Schreiber, 2008). Asterisks denote statistical significance (p-value < 0.05).

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in kea1kea2 mutants, and mirrored the recovery of the ionome shown in Table 1. Treating

kea1kea2 with NaCl also resulted in greater energy partitioning to ΦII, and less partitioning to

ΦNO (Fig. 3.3C-E). It should be noted that lower treatments of KCl (25 and 50 mM) also had a

negative effect on kea1kea2 biomass and photosynthesis (data not shown). Taken together, these

results indicate loss of KEA1/2 results in significant photodamage/photoinhibition, and relatively

less photochemistry. The fact that KCl exacerbates this phenotype while NaCl rescues it suggests

that K+ overaccumulation in the leaf impairs photosynthesis. As KEA1/2 are plastid envelope-

targeted, it is likely that excess K+ accumulates in the plastid stroma. Preliminary ionomics data

on isolated plastids shown in Chapter 2 (Fig. 2.7) revealed kea1kea2 plastids contained more K

than WT plastids. Na+ treatment may counteract K+ hyperaccumulation by competing for uptake

into the cell, or into the plastid. This is supported by the observation that treatment with exogenous

NaCl reduces K concentrations to WT levels in kea1kea2 (Table 3.1).

To ensure that K+ toxicity rather than Mg2+ deficiency was directly the cause of the

kea1kea2 phenotype, we also grew the kea1kea2 mutant on media with several concentrations of

MgCl2 (Supp. Fig. 3.1). We discovered that 75 mM MgCl2 is toxic to all genotypes (data not

shown). kea1kea2 treated with lower concentrations of MgCl2 (7.5 and 25 mM) displayed small

increases in Fv/Fm and ΦII, but these changes were only statistically significant for one independent

line. Based on this evidence, it seemed K+ toxicity was likely causal for the kea1kea2 phenotype,

and reduced leaf-levels of Mg were related to systematic decreases in chlorophyll content induced

by perturbation of photosynthesis. To better understand the molecular consequences caused by

hyperaccumulation of K+ in the plastid, we investigated nuclear gene expression in the mutant lines

compared to the WT under control conditions and NaCl treatment.

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Insights into the molecular consequences of disturbed chloroplast ion transport using nuclear

transcriptomics.

Aerial tissue from wild-type plants and the two independent T-DNA double mutant

kea1kea2 lines grown on control and NaCl

media as shown in Fig. 3.3 were harvested.

Subsequently, total mRNA was extracted,

converted into a cDNA library, and sequenced

on an Illumina platform. The standard Tuxedo

Suite software was used for transcript

alignment and differential expression analysis

(Trapnell et al., 2012). Each sample had high

coverage and sequence quality (Supp. Table

3.1). To ensure there were no systematic

errors from library preparation, we generated

a density plot of the FPKM expression values

for all samples (Supp. Fig. 3.2). The plot did not show any dramatic alteration in distribution for

any of the replicates, leading us to conclude no errors were introduced during library prep.

Principal Component Analysis (PCA) of gene expression values output from Cufflinks indicated

that most variance in gene expression results from sample identity, as sample replicates cluster

nicely in relation to the first principal component, which accounted for 69.5% of the variance (PC1,

Fig. 3.4). However, an additional factor appears to also contribute to variation in gene expression

in the samples, as indicated by the wide spread of samples in regard to the second principal

component, which accounted for 19.7% of the variance (PC2, for review of PCA analysis in

Fig. 3.4: PCA Analysis. Principal component analysis of normalized gene expression values output from Cufflinks. PCA was done on the online Illumina RNA-SEQ analysis platform Basespace™.

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sequencing data see Ringner 2008). Under control conditions, 3630 differentially expressed genes

(DEGs) were identified when comparing WT and both kea1kea2 double mutant lines (Table 3.2,

Supp. Fig. 3.3). Interestingly, when WT and kea1kea2 were compared under salt-stress conditions

which rescue development and photosynthesis in double mutants, the number of DEGs decreased

to only 1109 genes (Table 3.2, Supp. Fig. 3.3). The two lists of DEGs only shared 242 upregulated

genes, and 358 downregulated genes, suggesting that many of the genes that are deregulated in

mutant plants under control conditions are no longer differentially expressed compared to WT

after salt treatment (Supp. Fig. 3.4). According to the Arabidopsis protein subcellular localization

database tool SUBA4 MMAP (Hooper et al., 2017), there was an approximately 2-fold over-

representation of A. thaliana genes encoding chloroplast-localized proteins in the DEG list for the

control-treated WT vs. kea1kea2 (Fig. 3.5B) compared to the High Confidence Marker standard

(Suppl. 4A). For the DEG list comparing NaCl-treated WT with NaCl-treated kea1kea2 (Fig.

3.5C), there was a 3-fold overrepresentation of chloroplast-targeted genes compared to the HCM

standard (Fig. 3.5B). Overall, kea1kea2 disproportionately alters the expression of chloroplast-

targeted gene products under both treatments. Note that because of different lengths of input DEG

lists, comparisons of MMAP results between the WC_KC and WS_KS lists is not informative.

Table 3.2: Differentially Expressed Genes (DEGs)

Comparison Abbreviation Number of DEGs Up Down Total Col-0 control vs. kea1kea2 control WC_KC 1640 1990 3630 Col-0 NaCl vs. kea1kea2 NaCl WS_KS 525 584 1109 Col-0 control vs. Col-0 NaCl No further analysis 4130 4581 8711 kea1kea2 control vs. kea1kea2 NaCl No further analysis 3310 3316 6426 Col-0 control vs. kea1kea2 NaCl No further analysis 3544 4088 7632

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Functional enrichment analyses provide evidence that chloroplast gene expression is affected by

loss of KEA1/2

To deduce functional information from the nuclear transcriptomic dataset we employed the

functional enrichment tool FUNC-E (Ficklin and Poehlman, 2016). We used FUNC-E with three

Fig. 3.5: SUBA4 Multiple Marker Abundance Profiling (MMAP) tool applied to lists of Differentially Expressed Genes (DEGs) reveals many differentially expressed transcripts in kea1kea2 encode chloroplast-targeted proteins. (A) The High Confidence Marker set (HCM standard) used by SUBA4-MMAP shows the distribution of subcellular localizations for the A. thaliana proteome based on mass spectrometry. (B) SUBA4-MMAP was applied to the list of DEGs from the WT vs. kea1kea2 comparison under control conditions. (C) SUBA4-MMAP applied to the list of DEGs from the WT vs. kea1kea2 comparison under NaCl stress.

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independent annotation systems: functional annotations from the Gene Ontology (GO) database,

protein domains from InterPro, and protein families from PFAM. Briefly, the more DEGs assigned

to a particular annotation (i.e. GO term, PFAM family, InterPro domain), the lower the FDR-

adjusted Fisher p-value for this annotation (Fig. 3.6).

Supporting our previous finding that genes encoding chloroplast-targeted proteins were

overrepresented in our dataset (Fig. 3.5), we found most enriched GO terms when comparing

kea1kea2 to WT under control conditions (WC_KC) related to the broad categories of

chloroplast/photosynthesis, light signaling, and abiotic stress response. Protein domains and

families relating to all three of these categories were also enriched based on InterPro and PFAM

databases, corroborating the notion that loss of the inner envelope carriers KEA1 and KEA2

primarily impacts key processes linked to chloroplast function. Furthermore, we found enrichment

in six GO terms assigned to abiotic stress factors, namely cold, wounding, drought, salt, and

oxidative stress. Many of these annotations were also significantly enriched under NaCl, so were

unlikely to be causal for the kea1kea2 phenotype.

To gain insights into the delayed greening phenotype and the NaCl dependent rescue

mechanism in kea1kea2 mutants, we focused on annotations that were significantly enriched in

the WC_KC comparison, but not in the NaCl-treated WT-kea1kea2 comparison (WS_KS) that

were related to the plastid. This approach unveiled several GO terms which prompted further

investigations. Firstly, enrichment of the term “chlorophyll biosynthesis” (GO:0015995)

supported earlier results showing reduced chlorophyll levels in kea1kea2 and subsequent NaCl-

mediated recovery (Kunz et al., 2014). Additionally, several GO terms related to organellar gene

expression triggered our interest, as plastid gene expression (PGE) is a key process required for

chloroplast development (Bollenbach et al., 2005; Stoppel and Meurer, 2011; Tiller and Bock,

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Fig. 3.6: Functional enrichment analysis reveals that significantly Differentially Expressed Genes (DEGs) in kea1kea2 compared to the WT under control conditions are often associated with specific GO terms, INTERPRO domains and PFAM families. Some terms, domains and families are no longer significantly enriched when comparing kea1kea2 and WT when both are treated with NaCl. FUNC-E functional enrichment analysis of Differentially Expressed Gene (DEG) lists comparing the consensus of both kea1kea2 lines to WT under control (left column; WC_KC) and NaCl treatment (right column; WS_KS). The most significantly enriched annotations from the WC_KC are shown with corresponding results for the WS_KS comparison. GO terms, INTERPRO domains and PFAM families were used as the ontologies for analysis. Color scale of boxes corresponding to each term represent smaller False Discovery Rate (FDR) adjusted Fisher’s p-values (i.e., more significant = deeper blue). Grey boxes denote non-significantly enriched annotations (FDR-adjusted Fisher’s p-value ≥ 0.01). Color blocks over text correspond to a broad functional description of those annotations.

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2014). PGE-related terms included “nucleoid” (GO:0009295), “plastid chromosome”

(GO:0009508), and “chloroplast rRNA processing” (GO:1901259). Furthermore, analysis with

InterPro and PFAM databases revealed that nuclear-encoded proteins related to organellar gene

expression were enriched under the control treatment, but not under NaCl treatment. These

included organellar RNA-binding proteins such as pentatricopeptide repeat (PPR) proteins

(IPR002885, PF13812) and ribosome binding proteins (IPR003029, PF01926). This trend suggests

the delayed leaf greening and chloroplast development in kea1kea2 may be related to alterations

in chloroplast RNA processing, ribosome assembly or general chloroplast gene expression.

Notably, some other annotations which recovered after NaCl treatment related to other cell

structures (e.g., GO:0005773 vacuole, GO:0009505 plant-type cell wall) and non-plastid processes

such as phytohormone signaling (e.g., GO:0009734 Auxin-activated signaling pathway). We did

not further investigate genes related to these processes on the assumption they were pleiotropic

effects downstream of loss of plastid function.

When we took a closer look at the individual genes in respective functional GO annotations

(Fig. 3.7), it became apparent that transcripts were not always uniformly deregulated in one

direction. For instance, in the case of the annotation “chloroplast organization” (GO:0009658, Fig.

3.7A) we found that only about 20% of the transcripts were suppressed, and the remaining

transcripts increased in expression in the WC_KC comparison. Interestingly, two of the most

suppressed genes with this annotation encode the GOLDEN2-LIKE1 (GLK1) and GOLDEN2-

LIKE2 (GLK2) transcription factors (TFs). Together, GLK1/2 coordinate the expression of many

photosynthesis associated nuclear-encoded genes (PhANGs) and genes controlling chloroplast

development (Waters et al., 2008; Waters et al., 2009). Both TF mRNAs return to WT levels under

salt recovery. In the case of “chlorophyll biosynthesis” (GO:0015995, Fig. 3.7B), about 50% of

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Fig. 3.7: Log2 fold-change expression of individual genes that are significantly differentially expressed in kea1kea2 compared to WT under control conditions are no longer differentially expressed when comparing kea1kea2 and WT when both are treated with NaCl. (A-H) Log2 fold change gene expression for significantly differentially expressed genes (FDR-adjusted p-value < 0.05) from the WT vs. the two independent kea1kea2 T-DNA loss-of-function lines under control conditions (WC_KC1 and WC_KC2 for comparisons with kea1-1kea2-1 and kea1-2kea2-2 respectively) and corresponding information for the NaCl treatment (WS_KS1 and WS_KS2 for respective kea1kea2 lines). Red boxes denote statistically significant upregulation; blue boxes denote statistically significant downregulation. Grey boxes denote gene is not significantly differentially expressed (NS, FDR adjusted p-value ≥ 0.05).

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DEGs were suppressed, but the other half increased in expression. Notably, many downregulated

genes associated with this GO term encode proteins which catalyze or regulate committed steps in

the chlorophyll biosynthesis pathway during de-etiolation. These include HEMA1 (McCormac et

al., 2001), GUN4 (Adhikari et al., 2011), PORA (Paddock et al., 2012) and DXS (Mandel et al.,

1996; Estévez et al., 2001). The transcripts that increased in expression were generally not

involved in key chlorophyll biosynthesis reactions. Gene transcripts annotated with “de-etiolation”

(GO:0009704, Fig. 3.7C), were also primarily downregulated. In contrast, transcripts associated

with organellar gene expression related annotations (Ribosomal protein S1, IPR003029; “Plastid

Chromosome”, GO:009508; PPR Domain, PF13812; 50S Ribosome Binding GTPase, PF01926;

and “Plastid rRNA Processing”, GO:1901259) were almost universally significantly upregulated

in kea1kea2, although the degree of transcript increase varied (Fig. 3.7D-H). Strikingly, across all

annotations, we observed a reversion of most of these effects on gene expression in NaCl-stressed

plants, i.e., the transcription of two independent kea1kea2 mutants behaved more similarly to WT

under NaCl-stress conditions.

Additionally, even some annotations still significantly enriched under NaCl treatment in

the FUNC-E analysis displayed partial NaCl-mediated recovery of specific gene expression. These

annotations included light harvesting complex (GO:0030076, Fig. 3.8A), red and far-red light

signaling (GO:0010017, Fig. 3.8B) and circadian rhythm (GO:0007623, Fig. 3.8D). For each of

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these GO terms, most transcripts were downregulated under control treatment. However, NaCl

treatment returned expression to WT levels as described above. Independent of the FUNC-E

analysis, we hand-curated a list of TOC and TIC complex genes, which are responsible for plastid

protein import and therefore highly relevant for organelle biogenesis and plastome gene expression

Fig. 3.8: Log2 fold change expression of genes with annotations of interest. (A-D) Log2 fold change gene expression for significantly differentially expressed genes (FDR- adjusted p-value < 0.05) from the WT vs. the two independent kea1kea2 T-DNA loss-of-function lines under control conditions (WC_KC1 and WC_KC2 for comparisons with kea1-1kea2-1 and kea1-2kea2-2 respectively) and corresponding information for the NaCl-treatment (WS_KS1 and WS_KS2 for comparisons with kea1-1kea2-1 and kea1-2kea2-2 respectively). Red boxes denote statistically significant upregulation; blue boxes denote statistically significant downregulation. Grey boxes denote gene is not significantly differentially expressed (NS, FDR adjusted p-value ≥ 0.05).

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(PGE) (Jarvis and Soll, 2001; Köhler et al., 2016). Most transcripts in this category were

significantly upregulated in the mutant under control conditions (Fig. 3.8C).

In summary, the transcriptomics of nuclear genes further supported the NaCl-induced

phenotypic recovery reported for kea1kea2 double mutants (Kunz et al., 2014). More importantly,

the increased expression of PPR proteins, organelle RNA-binding proteins, and plastid ribosomal

proteins in kea1kea2 and their subsequent NaCl rescue suggests a link between ion transport and

PGE that may explain the delayed-greening phenotype in kea1kea2.

In collaboration with MPI-Golm: Simultaneous loss of KEA1 and KEA2 affects chloroplast rRNA

maturation

Intact PGE is a prerequisite for chloroplast development, integrity and function (Pogson and

Albrecht, 2011; Börner et al., 2015). The upregulation of plastid RNA and ribosome binding

proteins uncovered in the RNA sequencing (RNA-SEQ) experiment suggested that PGE might be

impacted in kea1kea2 loss-of-function mutants. Therefore, we worked with Dr. Reimo Zoschke’s

group at the Max Planck Institute of Molecular Plant Physiology in Golm to directly investigate if

disturbances in K+/H+ exchange across the chloroplast envelope caused by simultaneous loss of

KEA1/2 affect PGE on the transcriptional and post-transcriptional level. This was achieved by

analyzing chloroplast mRNA accumulation and translation by transcript and ribosome profiling

with a plastid microarray (Zoschke et al., 2013; Trösch et al., 2018). As in the NaCl-rescue

experiment described previously (Fig. 3.3), our collaborators compared chloroplast gene

expression from kea1-1kea2-1 and the WT under control and NaCl treatment (Fig. 3.9). They were

able to document some alterations of transcript accumulation (Fig. 3.9A), translation output

(footprint abundances, Fig. 3.9B), and translation efficiency (ribosome footprint abundances

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Fig. 3.9: Average relative translation output, transcript accumulation and translation efficiency of kea1kea2 compared to WT show no substantial specific changes in plastid gene expression under NaCl stress vs control conditions. However, kea1kea2 exhibits a plastid rRNA processing defect. (A) The ratio of transcript accumulation, (B) translation output (ribosome footprint abundances) and (C) translation efficiency (ribosome footprint abundances normalized to transcript accumulation) of all chloroplast reading frames is shown as log2 fold change ratio kea1-1kea2-1 vs. WT for both control (dark gray bars) and NaCl stress conditions (light gray bars). Positive or negative values depict a relative increase or decrease in expression in kea1-1kea2-1 relative to WT (average values of n = 2; gray dots represent individual biological replicates). The ocher-shaded areas indicate less than two-fold change. The dashed vertical lines separate genes according to the complexes their protein products reside in: Rub. = Rubisco, PSI = Photosystem I, PSII = Photosystem II, Cyt.b6f = Cytochrome b6f complex, ATP Syn. = ATP synthase, Poly. = RNA Polymerase, Others = other proteins. (D-G) Average signal intensity along length of (D) 16S, (E) 23S, (F) 4.5S, (G) 5S plastid rRNA transcripts in the WT and kea1-1kea2-1 under control and NaCl stress conditions. White bars represent signal from the 5’ and 3’ non-coding regions, and the gray bars represent signal from the coding portion of the rRNA transcript. The bars are the average values of n = 2; the black dots represent individual biological replicates.

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normalized to transcript abundances; Fig. 3.9C) in kea1kea2 compared to the WT. However, for

most genes the log2 fold change was low (less than 2; ocher-shaded areas in Fig. 3.9). More

importantly, not a single chloroplast gene exhibited substantially altered expression between

kea1kea2 and WT when comparing NaCl treatment and control conditions. Altogether, the mild

changes in the expression of specific chloroplast genes are an unlikely cause for the observed

phenotypic differences between NaCl and control conditions. This clearly contrasts our

observations on alterations in nuclear gene expression.

The probes on the chloroplast microarray also allow for semi-quantitative estimation of the

rRNA-processing status, although it should be kept in mind that saturation effects can affect these

estimations due to the exceptionally high abundance of mature rRNAs. However, in kea1kea2

control samples we observed unusually high signal intensities from probes covering 5’ and 3’ non-

coding regions of 16S (Fig. 3.9D), 23S (Fig. 3.9E), and 4.5S rRNAs (Fig. 3.9F). We did not

observe this effect for 5S rRNA (Fig. 3.9G). Typically, 5’ and 3’ regions give lower signals, as

seen in the WT, because they are cleaved during rRNA maturation (Manavski et al., 2018). NaCl

treatment resulted in reduction of signal from the non-coding regions and moderate increases in

signal from the coding regions in kea1kea2 compared to WT. Hence, our results indicate there is

disturbed rRNA maturation in kea1kea2.

kea1kea2 phenocopies mutants with 16S rRNA defects

In order to further investigate the potential rRNA processing effect, we compared the phenotype

of kea1-1kea2-1 to two previously described mutants: rap-1 (Kleinknecht et al., 2014) and rps5-2

(Fig. 3.10; Zhang et al., 2016). In both mutants, the loss of a nuclear-encoded RNA-binding protein

results in a chloroplast rRNA maturation defect. To confirm disruption of chloroplast rRNA

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function, we subjected all three mutants to increasing concentrations of spectinomycin, which

binds 16S rRNA and inhibits translation (Brink et al., 1994). All three mutants were more

susceptible to the inhibitor than the WT, indicating a preexisting deficit of functional 16S rRNA

(Fig. 3.10A). The rap-1 mutant was the most sensitive (survival rate < 50% at 1 µg ml-1), rps5-2

was the least sensitive (survival rate < 50% at 5 µg ml-1), and kea1-1kea2-1 had an intermediate

susceptibility (survival rate < 50% at 3 µg ml-1). When we grew kea1-1kea2- 1, rap-1 and rps5-2

Fig. 3.10: Loss-of-function lines with plastid 16S rRNA maturation defects resemble kea1kea2. (A) Percent survival rates for WT, kea1-1kea2-1, rap-1 and rps5-2 phenocopies on increasing concentrations of plastid translation inhibitor spectinomycin (± SEM, n = 9-10). Asterisks denote concentration where mean survival rate is significantly below WT survival (p-value < 0.05). (B) RGB image of WT, kea1-1kea2-1, and phenocopies rps5-2 and rap-1. (C) False color image of image of maximum quantum yield of PSII (Fv/Fm) for soil-grown plants. (D) Plot of Fv/Fm for ½ MS grown plants with or without NaCl (± SEM, n = 51-94). Different letters denote significantly different means (p-value < 0.05).

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side-by-side, all three mutants displayed a delayed-greening phenotype with chlorotic young

leaves, which improved with leaf age (Fig. 3.10B). We then investigated photosynthetic light

harvesting efficiency and found that the young leaves of all three mutants had significantly lower

Fv/Fm (Fig. 3.10C). Again, the phenotype was most extreme in rap-1, while rps5-2 was less

affected than the kea1-1kea2-1 mutant. Not surprisingly, kea1-1kea2-1 partially recovered in the

presence of NaCl, yet neither rap-1 nor rps5-2 showed a significant Fv/Fm increase, although a

slight, nonsignificant Fv/Fm increase was noticeable in rap-1 (Fig. 3.10D).

In collaboration with MPI-Golm: Northern blot further supports presence of rRNA processing

defects in kea1kea2.

Next, our collaborators carried out RNA gel blots to probe all four plastid rRNA species (Fig.

3.11). In the case of the 16S rRNA three different probes were hybridized: functional transcript,

5’, and 3’ extensions transcripts. For 23S, 4.5S, and 5S rRNAs the functional transcript regions

were probed (Fig. 3.11A). Substantiating the microarray results (Fig. 3.9), we found unprocessed

16S, 23S, and 4.5S rRNA transcripts to accumulate in kea1kea2 under control conditions (Fig.

3.11B). A similar, yet more severe maturation defect was found in control rap-1 and rps5-2 plants

(Fig. 3.11B). Interestingly, NaCl treatment reduced the accumulation of immature 16S, 23S, and

4.5S RNAs in kea1kea2 (Fig. 3.11B-C) but had no effect on rRNA maturation in rap-1 or rps5-2

(Fig. 3.11B-C). The maturation of 5S rRNA transcripts was similar across genotypes and

regardless of the treatment. In summary, the virescent phenotype in kea1kea2 correlates with

rRNA processing defects, both which are rescued by NaCl exposure. Therefore, NaCl-mediated

rescue of the kea1kea2 phenotype correlates with rescue of rRNA processing.

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Fig. 3.11: Plastid rRNA processing defects in the kea1kea2 mutant are alleviated under NaCl stress conditions compared to control conditions. (A) Physical map of the plastid rRNA transcript with binding position of probes (black lines) used in northern blots. (B) From top to bottom: Northern blot analyses of 16S rRNA accumulation using probes against the 5’ non-coding region (5´), the 16S rRNA coding region (16S) and the 3’ non-coding region (3´). In the middle blot, the upper and lower bands represent unprocessed and mature 16S rRNA, respectively. (C) The same Northern blots from (B) rehybridized to analyze other plastid rRNAs. From top to bottom: 23S, 4.5S, and 5S rRNA accumulation. Sizes of marker bands are indicated on the left side in kilonucleotides (knt). Pictures of methylene blue-stained chloroplast and cytosolic rRNAs (M.B.) are provided below each blot to illustrate equal loading. Black asterisks show the rRNA precursor bands. Samples analyzed include control and salt-treated WT, kea1-1kea2-1, and two lines with previously characterized 16S maturation defects, rap-1 and rps5-2.

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Steady-state levels of several plastome-encoded proteins are decreased in kea1kea2 mutants.

To investigate how disruption of rRNA metabolism (as demonstrated by transcriptomics and

northern blots) may impact chloroplast protein level at different leaf developmental stages, we

compared steady-state levels of nuclear- and plastome-encoded proteins in kea1kea2 young pale

leaves and mature green leaves with respective WT tissues of the same developmental stage (Fig.

3.12, Supp. Fig. 3.5). Initially, we checked protein levels of the vacuolar–ATPase Epsilon subunit

as a loading control. V-ATPase amounts were unchanged between the genotypes and leaf ages.

Furthermore, RNA-SEQ data also showed that V-ATPase was not differentially expressed

between the genotypes (AT4G11150). KEA1 and KEA2 carriers were slightly more abundant in

young WT leaves, but as expected were absent from kea1kea2 protein extracts. Interestingly,

plastome-encoded proteins RbcL and PsbA revealed lower abundance in young pale kea1kea2 leaf

tissue compared to young WT tissue. For RbcL the same result was also visible in the Coomassie

Blue gel stain. However, RbcL was detectable at WT levels in the older green leaves of kea1kea2.

The small nuclear-encoded subunit of the Rubisco complex (RbcS), which is sometimes

considered a Photosynthesis Associated Nuclear-Encoded Gene (PhANG) (Ruckle et al., 2007;

Hills et al., 2015), varied from being unchanged in all samples (Fig. 3.12) to decreased specifically

in kea1kea2 young leaves (Supp. Fig. 3.5). Lhcb1, the only other PhANG we were able to probe

by immunoblotting, was markedly reduced in young kea1kea2 leaves but present at WT level in

older tissue. Finally, the plastid protein import machinery subunits Tic110 and Tic40 were slightly

more abundant in kea1kea2 independent of the leaf age.

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Pulse-chase studies reveal altered translation in kea1kea2 chloroplasts.

The ribosome profiling method can readily detect translation defects for specific chloroplast genes.

However, the rRNA processing defect in kea1kea2 may cause a general chloroplast translation

decrease that may have gone undetected in our ribosome profiling as indicated by altered steady-

state levels of plastome-encoded proteins (Fig. 3.12, Supp. Fig. 3.5). Hence, we performed pulse-

chase assays to determine if in kea1kea2 the overall synthesis or turnover of plastome-encoded

proteins was altered due to translational defects. In consideration of the characteristic delayed-

greening phenotype in kea1kea2, we again investigated young and mature leaves from both

genotypes separately (Fig. 3.13, Supp. Fig. 3.6). Initially, we analyzed total chloroplast translation

Fig. 3.12: Immunoblots reveal lower steady-state levels of plastid-encoded proteins. Immunoblots for steady-state levels of plastid and nuclear-encoded proteins in young (Y) vs. mature (M) leaf tissue from WT and kea1kea2. Plastid-encoded proteins include Rubisco large subunit (RbcL) and the D1 reaction center of photosystem II (PsbA). Nuclear-encoded plastid-targeted proteins include Rubisco small subunit (RbcS), light harvesting chlorophyll a/b binding protein (LHCb1), protein import complex components Tic110 and Tic40, and K+/H+

antiporters KEA1/2. Vacuolar ATPase (V-ATPase) was used to probe the abundance of non-chloroplast proteins and demonstrate equal loading of samples. Coomassie stain of gel is aligned below blots to show relative amounts of total protein loaded. Additional blot shown in Supp. Fig 3.5.

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and protein turnover rates under illumination in young and mature leaf samples from the WT and

the mutant. The pulse portion of the experiment revealed significantly lower rates of 35S

Fig. 3.13: Pulse-chase experiments reveal decreased translation rates in kea1kea2. (A) Representative autoradiograph from the pulse-chase analyses. Leaf discs from young or mature WT and kea1kea2 leaves were harvested after 15 and 40 minutes during the pulse portion of the experiment (black labeled lanes). After chase with non-radioactive amino acids, samples were taken every 2 hours for a total of 400 minutes (red labeled lanes). Bands corresponding to rubisco large subunit (RbcL) and the D1 reaction center of PSII (PsbA) are clearly visible. Additional autoradiographs in Supp. Fig. 3.6 (B) Quantification of total protein produced in pulse-chase experiment (top), RbcL production (middle) and PsbA (bottom). Leaf discs were harvested after 0, 15, and 40 minutes during the pulse portion of the experiment. After chase with non-radioactive amino acids, samples were taken every 2 hours for a total of 400 minutes. Red line represents transition from pulse to chase stage. Total protein production was measured in a scintillation counter to quantify counts per minute (CPM). For RbcL and PsbA, intensity of autoradiograph was quantified in ImageJ and normalized to the intensity of the WT at the 40-minute timepoint (± SEM, n = 3). Green arrows adjacent to line indicate kea1kea2 has a significantly lower protein synthesis rate from the WT at given timepoint (p-value < 0.05).

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incorporation in kea1kea2 chloroplast proteins compared to WT specifically in young leaves (Fig.

3.13B). Protein degradation rates did not vary significantly over the course of the 320-minute chase

period. RbcL and PsbA, the two most abundantly translated proteins in chloroplasts, were utilized

to track the translation-degradation kinetic of individual proteins. Unexpectedly, RbcL translation

occurred to the same degree in young WT and kea1kea2 leaf tissue, yet older green kea1kea2

leaves showed a significant increase of RbcL translation compared to its respective WT control

tissue. RbcL protein was also very stable and no significant loss in labeling was detected in either

genotype during the chase portion of the assay. In contrast, PsbA translation rates were strongly

reduced in young pale kea1kea2 leaves. The difference between the genotypes was less dramatic

in older leaves. As expected, PsbA turnover was quick and took place at a similar rate in WT and

kea1kea2 during the 320-minute-long chase period. We conclude that overall chloroplast protein

production rates are lower in kea1kea2. This effect is most dramatic in the pale young mutant leaf

tissue.

Loss of KEA1/2 suppresses var2 phenotype in young leaves

It is known that drugs and mutations which compromise plastid rRNA processing, translation and

other aspects of PGE rescue the variegated phenotype of the chloroplast protease mutant var2

(Miura et al., 2007; Liu et al., 2010; Yu et al., 2011). The var2 mutants have decreased levels of

the plastid metalloprotease FTSH2, which plays a key role in PSII repair and prevents

photoinhibition by degrading damaged D1 reaction centers (reviewed in Kato and Sakamoto,

2018). Thus, variegated regions in var2 leaves are thought to be areas where excess photoinhibition

has impaired chloroplast development, resulting in white sectors containing abnormal plastids

(reviewed in Putarjunan et al., 2013; Kato and Sakamoto, 2018). Why second-site mutations in

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PGE-related genes specifically act as

suppressors is still unclear (Liu et al.,

2010; Putarjunan et al., 2013). Yet,

this suppression phenomenon has

been frequently employed as a tool

to determine if PGE is decreased in a

mutant of interest by introgressing

the respective mutant into the var2

background, (reviewed in Yu et al.,

2008; Putarjunan et al., 2013). We

isolated two independent

kea1kea2var2 triple mutant lines and found that the loss of KEA1/2 affects chloroplast gene

expression sufficiently to suppress the variegated var2 phenotype (Fig. 3.14). However, this

suppression was most effective in young leaves. In some triple mutant individuals, older leaves

did develop the characteristic white sectors, although to a smaller degree and less frequently than

var2 single mutants. This result supports our earlier finding that kea1kea2 mutants possess an

rRNA processing defect, like many other var2 suppressor mutants. Notably, the suppression in

young leaves coincides with decreased synthesis of PsbA based on our pulse-chase analysis (Fig.

3.13) and decreased steady state levels of PsbA and Lhcb1 (Fig. 3.12).

Collaboration with Soll Lab at LMU-Plastid protein import is not compromised in kea1kea2

mutants. Nuclear-encoded transcripts for plastid-targeted proteins are translated in the cytosol into

precursor proteins, which are then imported into the plastid stroma via translocation complexes

Translocon Outer Membrane Complex (TOC) and Translocon Inner Membrane Complex (TIC)

Fig. 3.14: Loss of KEA1/2 suppresses leaf variegation in var2-5. RGB image of WT, the two independent kea1kea2 loss-of-function lines, var2-5, and the two kea1kea2 lines introgressed into the var2-5 background. Note the presence of white patches on the old, but not the young, leaves of kea1kea2var2 mutants. kea1kea2 double mutants do not display any white patches.

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(Thomson et al., 2020). Since most chloroplast RNA-binding proteins are encoded in the nucleus,

proper chloroplast RNA processing requires the TOC/TIC import pathway (Zoschke and Bock,

Fig. 3.15: Chloroplast protein import rates are similar in kea1kea2 vs WT, indicating that import of RNA binding proteins is not impaired. (A) Autoradiograph (top) and Coomassie stain (bottom) of gels showing import of precursor proteins ferrodoxin reductase (FNR; left) and pyruvate dehydrogenase E1α subunit (PDH E1α; right) into isolated chloroplasts from WT and the two independent kea1kea2 lines. Top bands on autoradiograph (40 kDa for FNR and 47 kDa for PDH E1α) correspond to preprotein prior to import and removal of plastid transit peptide. Lower bands (35 kDa for FNR and 40 kDa for PDH E1α) correspond to the imported, processed protein. The third band for PDH E1α at 39 kDa is likely a late start artifact of the in-vitro translation system. (B) Quantification of imported proteins. The intensity of bands corresponding to imported FNR and PDH E1α were normalized to the intensity of all protein bands on the Coomassie stained gel excluding RbcL and the LHC proteins. These values were then normalized to the mean of the WT, resulting in a value for relative import as a % of import in the WT (± SEM, n = 3-14). Differing letters above the bars denote significantly different means (p-value < 0.05).

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2018). Therefore, we tested the protein import capacity of isolated intact kea1kea2 and WT

chloroplasts using the two well-established import substrates, ferredoxin-NADP+ reductase (FNR)

and pyruvate dehydrogenase E1 alpha subunit (E1α) to account for the posited translocon dual

substrate selectivity (Ivanova et al., 2004; Inoue et al., 2010). The data show that the kea1kea2

mutant plastids had protein uptake rates comparable to WT or even slightly higher (Fig. 3.15).

Therefore, the rRNA processing defect documented in kea1kea2 is not caused by compromised

plastid protein import resulting in a shortage of nuclear-encoded RNA-binding proteins in the

stroma.

DISCUSSION

Summary

In this study, we characterized the consequences from a loss of the two important chloroplast

K+/H+ exchangers KEA1/2. This was achieved through a systems biology investigation of

kea1kea2 loss-of-function lines. Earlier studies revealed that kea1kea2 double mutants are

characterized by major impairment of photosynthesis and changes in the leaf ionome dominated

by K+ (Kunz et al., 2014; Höhner et al., 2016b). We show these changes wcan be ameliorated by

exogenous NaCl treatment, but not by KCl treatment (Table 1, Fig. 3.3). Previous work also

revealed that kea1kea2 double mutants exhibit a delayed greening phenotype in young leaves with

altered chloroplast biogenesis (Aranda-Sicilia et al., 2016). In this study, we have collected strong

evidence that impairment of chloroplast development and delayed leaf greening in kea1kea2 likely

originate from a partial impairment of plastid gene expression (PGE).

We began our investigation by using RNA-SEQ to gain clues as to which regulatory and

metabolic pathways were most altered by loss of KEA1/2, yet recovered under NaCl treatment, a

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stress condition known to revert developmental defects in kea1kea2 (Kunz et al., 2014). Functional

enrichment analysis revealed numerous genes with photosynthesis and chloroplast-related

annotations were deregulated in kea1kea2. However, only a small set of these annotation groups

recovered to wildtype levels under NaCl treatment. Most of these annotations were related to

chloroplast RNA metabolism. A deeper investigation of these genes revealed that many PPR

proteins involved in chloroplast RNA processing and editing were upregulated in kea1kea2,

suggesting that a cellular program was triggered to compensate for a defect in chloroplast RNA

metabolism or other impairment of PGE. This prompted us to investigate plastid transcriptomics

and RNA metabolism directly using a plastid-targeted transcriptome and ribosome profiling

method.

Plastid transcriptomics and northern blotting indicated maturation defects for all

chloroplast rRNAs (16S, 23S 4.5S and 5S) resulting from loss of plastid ion transporters. This

defect may impact the function or assembly of chloroplast ribosomes, supported by pulse–chase

experiments showing hampered protein translation in kea1kea2 chloroplasts. The pulse-chase

experiment showed that in young leaves, the translation rate for chloroplast-encoded proteins was

impaired, in addition to a dramatic reduction specifically for PsbA, the D1 core subunit of PSII.

The specificity of this effect on young leaves further supports the notion that the ‘virescent’

phenotype (young pale leaves) is indeed a signature phenotype for mutants with compromised

PGE as posited before (Zhou et al., 2009; Zheng et al., 2016). Our data suggest that kea1kea2 null

mutants can be considered PGE-deficient mutants as well. Further support for this idea was

gathered from the established virescent mutants rap and rps5, both with plastid rRNA maturation

defects, which phenotypically resemble kea1kea2 mutants.

Additional evidence for disrupted PGE in kea1kea2 was documented in this study. Firstly,

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steady state levels of plastome-encoded proteins (RbcL, PsbA) were reduced especially in young

kea1kea2 leaf tissue. Nuclear-encoded plastid proteins were heterogeneously influenced by loss of

KEA1/2. Some proteins were upregulated in kea1kea2 young leaves (TOC, TIC), while other

proteins known to be the targets of stress-mediated retrograde signaling (LHCb) were

downregulated. Secondly, kea1kea2 mutants are more susceptible to spectinomycin, which

specifically inhibits plastid translation by binding 16S rRNA (Mohan et al., 2014). The increased

drug sensitivity in the kea1kea2 mutant could occur due to the 16S rRNA maturation defect in the

mutant reducing the activity or quantity of ribosomes, or independent effects on mature ribosome

activity due to loss of ion homeostasis. Finally, simultaneous loss of both envelope KEA carriers

partially rescues the var2 mutant phenotype. The leaf variegation phenotype of var2 mutants

originates from defective D1 protein turnover (Chen et al., 2000b; Lindahl et al., 2000). Ample

second-site genetic screens have manifested that gene mutations that suppress the leaf variegation

in var2 are almost exclusively tied to decreased PGE rates (Liu et al., 2010). Therefore, the var2

suppression has emerged as a genetic assay to verify the status of PGE in a given mutant

(Putarjunan et al., 2013). Intriguingly, the suppression of variegation in kea1kea2 was

heterogeneous, with complete suppression in young leaves, and older leaves exhibiting some

variegation. Authors have hypothesized that the mechanism of suppression is that general

reduction in PGE results in decreased ROS production, either through decreasing synthesis of the

PsbA protein (Putarjunan et al., 2013) or inducing retrograde signaling to suppress the expression

of Lhc components (Yu et al., 2008). Thus, reduced steady state levels of PsbA and Lhcb1 (Fig.

3.12) or decreased PsbA synthesis (Fig 3.13) specifically in kea1kea2 young leaves could explain

why the suppression of variegation is restricted to these tissues.

Ion homeostasis and plastid gene expression: a new hypothesis.

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Our findings raise the question why plastid gene expression is affected by the loss of KEA1/2

antiporters. Specifically, there are two open questions: 1) Is the primary effect from loss of KEA1/2

transporters perturbation of stromal ion homeostasis? 2) Is loss of stromal ion homeostasis the

direct cause of the rRNA processing and translation effects we quantified in kea1kea2? We use

our results in combination with evidence from the literature to build the case for a new hypothesis

linking ion homeostasis and PGE. We propose that altered stromal cation concentrations resulting

from loss of KEA1/2 impairs the interaction of nuclear-encoded RNA processing proteins with

target RNAs and/or impairs ribosomal assembly, thus inhibiting plastid translation.

KEA1/2 are likely a valve for K+ out of the stroma in light-exposed chloroplasts.

Direct transport assays with KEA protein reconstituted in liposomes and complementation

experiments employing K+ transporter deficient yeast and E. coli strains have confirmed K+/H+

exchange for all KEA family members (Aranda-Sicilia et al., 2012; Zhu et al., 2018; Tsujii et al.,

2019). The directionality of transport through KEA1/2 is likely determined by the prevailing K+

and H+ gradients across the plastid envelope. K+ concentrations in the cytosol (Leigh and Jones,

1984) and the plastid (Demmig and Gimmler, 1983; Robinson and Downton, 1984; Schröppel-

Meier and Kaiser, 1988) are thought to be roughly equivalent. However, precise stromal K

concentrations have not been quantified. The cytosol has a near-neutral pH (~7, Demes et al.,

2020), while the pH of the stroma fluctuates from near neutral in the dark to basic (~8) in a

photosynthesizing chloroplast (Su and Lai, 2017; reviewed in Höhner et al., 2016a). Therefore, pH

gradients favor K+ efflux in exchange from the stroma in exchange for protons in the light. In the

dark, the KEA antiporters could work the opposite direction. Indeed, a recent study showed that

loss of KEA1/2 prevents K+ -mediated pH changes in the stroma of isolated chloroplasts in the

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dark (Aranda Sicilia et al., 2021). We argue that loss of light-driven K+ efflux is the causal effect

for the developmental and photosynthetic phenotype in kea1kea2. While there is no published

data on the A. thaliana chloroplast ionome, our preliminary results from the previous chapter show

increased K in the kea1kea2 chloroplast, suggesting that loss of these two transporters prevents K+

efflux from the stroma. This is consistent with previously published data that kea1kea2

accumulates excess K in its leaves (Höhner et al., 2016b, Höhner et al., 2019), and our observation

that treatment with exogenous K+ exacerbates the kea1kea2 phenotype (Fig. 3.3). Thus, it is likely

that the primary effect of losing KEA1/2 is an accumulation of K+ in the stroma, accompanied by

linked changes in other ions. This is significant because in vitro studies by others suggest that

unbalanced stromal ion homeostasis may either inhibit the catalytic function of nuclear-encoded

chloroplast RNA-binding proteins or prevent binding to their respective target.

Altered concentrations of K+ and other cations impair protein-RNA interactions and ribosome

assembly in-vitro.

As stated before, the posttranscriptional and translational stages of plastid gene expression rely

heavily on the activity of RNA-binding proteins, including PPRs, ribosomal proteins, and

chloroplast ribonucleoproteins (Tillich et al., 2010; Zoschke et al., 2011). Our in-vitro import

assays suggested that protein uptake in kea1kea2 mutant chloroplasts proceeds at or above WT

rates. Furthermore, our RNA-SEQ data revealed a striking increase in many nuclear-encoded

chloroplast RNA-binding protein transcripts. Therefore, we conclude that a shortage of RNA-

binding proteins is not the cause of the rRNA processing defect in kea1kea2. Thus, reduced activity

or binding of these proteins to target RNAs due to stromal buffer disruptions may be to blame.

For instance, establishing PPR binding to target RNAs is strongly dependent on electrostatic

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forces. Even weak changes to RNA structure, such as secondary structures induced by NaCl,

disturb the binding site and protein-RNA interaction (McDermott et al., 2018). Furthermore,

optimal ribosomal function requires balanced concentrations of Mg2+, K+, and cationic polyamines

(Nierhaus, 2014). While it has been known for years that a lack or excess of K+ prevents proper

integrity and function of bacterial-type ribosomes (Bhaya and Jagendorf, 1984), research has

focused primarily on mechanistically understanding the importance of Mg2+ for prokaryotic

translation. Mg2+ ions play a key role in the stabilization of rRNA-protein interactions, ribosomal

assembly, and tRNA-ribosome binding (Horlitz and Klaff, 2000; Blaha et al., 2002; Hirokawa et

al., 2002; Konevega et al., 2004; Petrov et al., 2012). However, excess K+ outcompetes Mg2+ for

binding sites on ribosomal subunits, which impairs ribosome assembly and consequently protein

translation (Nierhaus, 2014). At this point, we cannot assign the phenotypic malfunction in

kea1kea2 specifically to stromal K+ level as we have not definitively quantified all of the elements

in chloroplast (i.e. Mg) due to the limitations of the TX-RF method. However, it is logical to

postulate the observed effects on plastid rRNA maturation and PGE are at least partially caused

by an increase in stromal K+ level in kea1kea2 mutants.

Further support for the plastid K+ imbalance hypothesis comes from the fact that NaCl

treatments rescue the rRNA maturation defects (and subsequently photosynthesis) in kea1kea2 but

not in RNA-binding protein deficient mutants rap and rps5. It is widely known that NaCl stress

results in K+ depletion because Na+ and K+ ions compete for root uptake (Munns and Tester, 2008;

Deinlein et al., 2014). Depending on the NaCl stress severity, leaf cells become K+ starved (Zhu

et al., 1998; Stepien and Johnson, 2009; Chao et al., 2013). This situation is favorable for kea1kea2

as less K+ can build up in the mutant chloroplasts, partially resetting the stromal ion homeostasis.

Indeed, ionomics data on leaf tissue shown in Table 1 indicates NaCl treatment specifically

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reduces K concentrations in kea1kea2 to WT levels. This effect explains the recovery of rRNA

maturation in kea1kea2 under NaCl treatment. A similar rescue effect is not possible if rRNA

maturation is directly hampered through the loss of an RNA-binding protein as exemplified in rap

and rps5 mutants. Interestingly, a recent study on the envelope Mn2+ transporter CMT1 also found

evidence for disrupted PGE in response to loss of ion homeostasis. However, in cmt1-1 mutants

the lack of stromal Mn2+ ions specifically decreased transcription rates and only as a consequence,

translation (Eisenhut et al., 2018).

Future directions

Our results reveal a previously undocumented influence of plastid ion transport and homeostasis

on organellar rRNA metabolism and translation. Furthermore, loss of ion homeostasis may

influence ribosomal assembly and activity independently of rRNA processing. Yet, how ion

transport and homeostasis mechanistically effect RNA metabolism and ribosome activity in vivo

remains to be elucidated in more detail. Future experiments to test chloroplast ribosome activity

and quantify overall protein-RNA interactions in the kea1kea2 background are necessary for a

definitive understanding. One such experiment would be to carry out ribosome loading analysis

in the kea1kea2 background. This technique uses a sucrose gradient in combination with RNA

blotting to quantify ribosome association with specific transcripts (Bock and Zoschke, 2018). This

type of analysis would provide additional evidence that ribosomal activity is decreased in

kea1kea2. To directly test RNA-protein interactions in the kea1kea2 background compared to WT,

we could carry out Electrophoretic Mobility Shift Assays (EMSA), in both genotypes using the

16S RNA precursor transcript as our target RNA. EMSA involves incubation of cell or organelle

lysate with a radioactive probe for a RNA molecule of interest, and subsequently detecting RNA-

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protein interactions based on size shifts in an RNA blot (Holden and Tacon, 2011). Alternatively,

FRET-style analyses using an assortment of GFP-tagged RNA-binding proteins of interest and

fluorescent nucleic acid dye Syntox orange can be used to quantify protein-RNA interactors where

potential protein interaction partners are known (Camborde et al., 2017). In the case of kea1kea2,

16S RNA interactions with previously characterized 16S rRNA-binding proteins such as PPR-type

protein RAP (Kleinknecht et al., 2014) or 16S-binding ribonuclease YbeY (Liu et al., 2015) could

be investigated in-vivo.

Alternative hypotheses.

Over the years, several alternative explanations for our results have been suggested. One such

explanation could be that KEA antiporters have an uncharacterized RNA-binding function. This

seems unlikely, as the KEA proteins do not contain any domains annotated with nucleic-acid

binding functions other than the NADPH/NADH binding capability of the C-terminal KTN

domain (Roosild et al., 2009). However, we could test this possibility by doing RNAse protection

assays followed by immunoblotting with an antibody specific to KEA1/2. This method has been

used previously to test RNA binding capabilities in proteins (Schmid et al., 2019).

Another plausible explanation is that the perturbation of photosynthesis in kea1kea2

mutants results in ROS accumulation, which has been shown to impair plastid translation by

inhibiting elongation factor G (EF-G, Nishiyama et al., 2011). This effect was initially described

in regard to the slowing of PsbA turnover in cyanobacteria in the presence of H2O2 and 1O2.

However, it is unlikely that ROS-mediated inhibition of EF-G is exclusively the cause of reduced

PGE in kea1kea2 for several reasons. First, while ROS inhibition of EF-G could reduce translation

rates in kea1kea2, ROS have never been shown to interfere with plastid rRNA processing or induce

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rRNA defects, a phenomenon we clearly detected in the kea1kea2. Second, the chase portion of

the experiment revealed similar rates of PsbA turnover in the WT and kea1kea2, indicating that

kea1kea2 is not suffering from irreparable damage to PSII due to turnover defects (Fig. 3.13b).

Third, pulse-chase analysis carried out at low light intensities of ~10 µE also showed decreased

protein biosynthesis in kea1kea2 compared to the WT (Supp. Fig. 3.7). If the decrease in

translation rates exhibited by kea1kea2 were the result of ROS-mediated inhibition of EF-G, one

would expect that protein synthesis should be similar in kea1kea2 compared to the WT in light

conditions that minimize oxidative stress.

CONCLUSION

This work has shown in-vivo that KEA1/2-dependent K+/H+ transport is essential for ribosomal

maturation or assembly, and thus is a key factor for optimal plastid gene expression. Based on our

Fig. 3.16: Model depicting the influence of chloroplast ion homeostasis on Plastome Gene Expression (PGE) in WT plants, and in the kea1kea2 mutant under control and salt-treatment conditions. (A) WT cell; (B) kea1kea2 cell; (C) kea1kea2 cell with exogenous NaCl treatment. Abbreviations are as follows = Plastome Gene Expression; cRBPs = chloroplast RNA-Binding Proteins.

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results, we built a speculative model depicting how loss of KEA1/2 transporters influences PGE

and plastid development. In a WT leaf cell (Fig. 3.16A), KEA1 and KEA2 transporters in the

plastid envelope operate to maintain ion homeostasis in the stroma as demonstrated in chapter 2.

With proper levels of stromal ions, chloroplast RNA binding proteins (cRBPs) can properly bind

and mediate processing of plastid RNAs. This allows for maturation of rRNA, and subsequently

the assembly of functional ribosomes which can catalyze protein biosynthesis. PGE progresses as

usual in a WT leaf cell, resulting in timely chloroplast development. In a kea1kea2 leaf cell (Fig.

3.16B), loss of the two K+/H+ antiporters perturb plastid ion homeostasis. The stromal buffer

conditions are not ideal for cRBP interaction with RNA targets, and there may be additional effects

on ribosomal assembly and function. This results in a lower translation rate, and overall lower rates

of PGE. Hence, plastid biogenesis is delayed. In the leaf cell of a kea1kea2 mutant treated with

exogenous NaCl, stromal ion homeostasis is partially rebalanced, perhaps due to reduced uptake

of K. The ion content of the plastid is better suited for protein-RNA interactions and ribosomal

assembly/activity. Therefore, PGE and subsequently plastid development is partially rescued.

These results will inform further studies related to plastid development and photosynthesis

bioengineering by underscoring the importance of maintaining homeostasis of inorganic

components, i.e., the chloroplast stromal buffer. In the future, we will use this work as a basis to

do direct studies of RNA-protein interactions in the stroma of ion transporter mutants like

kea1kea2.

METHODS

Germplasm and General Plant growth. Homozygous kea1-1kea2-1 and kea1-2kea2-2 were

obtained from a previous study (Kunz et al., 2014). The var2-5 point mutant was previously

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published (Chen et al., 2000a). All other loss-of-function lines were obtained from ABRC. Lines

were confirmed to be homozygous using PCR with one gene-specific primer and one T-DNA

specific primer. For a full list of lines and primers see Appendix B. Unless otherwise specified,

plants were grown as follows; seeds where surface sterilized in 70% ethanol, then spread on

sucrose-free ½ MS agar media (Murashige and Skoog, 1962) and vernalized for 48 hours in the

dark at 4° C. Seed were then placed in a Percival Growth Chamber with 150 µmol photons m-2s-1

of light, 16 h: 8 h light: dark cycle. Growth temperatures were 22° C in the light and 18° C in the

dark. Plants were grown for 7 days then either transferred to soil or to treatment plates.

Ionomics on NaCl-rescued WT and kea1kea2 lines. For ionomics, unpublished data from Ricarda

Höhner was reanalyzed. Plant growth and ionomics were conducted as described in Höhner et al

2016b. In brief, plants were grown on soil for 4 weeks, and watered with either ordinary ½ MS

media (control) or with ½ MS containing 75 mM NaCl (NaCl). 1 leaf disc was harvested from

each true leaf (leaves 1-4) using a tissue punch. Discs were dried at 80° C for 48 hours and

weighed. The discs were then digested in nitric acid and measured on a Picofox TX-RF instrument

(Bruker, Germany) as described in Höhner et al., 2016b.

For analysis, the values for the tissue free blank were subtracted from the sample values.

Then, sample values were normalized to dry wieght. The normalized values for all for leaf punches

were averaged. Statistical differences in mean K content were then determined by a two-way anova

and and a series of pairwise Tukey’s multiple comparison tests (p-value < 0.05).

Plant Growth for PAM and RNA-SEQ. Seeds from Col-0 wildtype (WT) A. thaliana and two

independent kea1kea2 double mutant lines were prepared as described in the general plant growth

section. One-week-old seedlings were then transferred to ½ MS agar media with or without 67.5

100

mM NaCl and grown for an additional 15 days in the conditions described above (plants were 22

days old at harvest).

PAM photosynthesis measurements. Chlorophyll fluorescence measurements were taken as

previously described (Kunz et al., 2009). Plants were dark adapted for 20 minutes prior to imaging.

Imaging was carried out on an Imaging PAM M-series chlorophyll fluorometer (Walz) using a

built-in induction protocol for chlorophyll fluorescence kinetics with a photosynthetically active

radiation (PAR) intensity of 185 photons m-2s-1.

RNA Isolation and Sequencing. For each treatment and genotype, there were 3 biological

replicates. Total RNA was isolated from leaf tissue using the GeneJet Plant RNA purification kit

(ThermoFisher). Contaminating genomic DNA was removed via treatment with DNAse I

(ThermoFisher). Total RNA was then converted into a cDNA library using an OligoDT polyA tail

pulldown kit. Sequencing was carried out on an Illumina HiSeq 2500 at the Genomics Center at

Washington State University, Spokane. Reads were 100 base pairs long, paired end, with over 15

million reads per sample. For details, see Supp. Table 1. Raw sequencing data was reposited at

the NCBI SRA public database (accession PRJNA573960,

https://www.ncbi.nlm.nih.gov/sra/PRJNA573960).

RNA Sequencing (RNA-SEQ) Data Analysis. The raw fastq files were uploaded to the Illumina

platform Basespace for further analysis through the Tuxedo Suite of RNA-SEQ analysis software.

This pipeline includes the programs Bowtie, TopHat, Cufflinks, and Cuffdiff (Trapnell et al.,

2012). These programs aligned raw reads to the TAIR10 Arabidopsis reference genome (Berardini

et al., 2015), assembled transcripts, and subsequently called differentially expressed genes based

on pairwise comparisons between each sample group. For this analysis, the False Discovery Rate

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(FDR) adjusted p-value (q-value) was set to 0.05 to call significantly Differentially Expressed

Genes (DEGs). Subsequently, these genes were matched with a functional annotation from

TAIR10 (Yon Rhee et al., 2003). The lists of DEGs from the WT vs. mutant comparisons under

both control and NaCl conditions were then further filtered for genes that were differentially

expressed in both independent kea1kea2 mutant lines in the same direction to form a consensus

DEG list. In other words, each gene on the consensus list was significantly up- or downregulated

in both the WT comparison with kea1-1kea2-1 and the comparison with kea1-2kea2-2. Values

from the consensus list for the control and NaCl treatment are labeled as “WC_KC” and WS_KS

respectively in the figures and text. The consensus lists for both treatments were first analyzed by

the subcellular compartment localization prediction tool SUBA4 (Suppl. Fig. 4, (Hooper et al.,

2017)). The consensus lists for both treatments were then used for functional enrichment analysis

through the tool FUNC-E to find significantly enriched Gene Ontology (GO) terms, PFAM protein

families, and INTERPRO domains in kea1kea2 compared to WT under control and NaCl

conditions (Ficklin and Poehlman, 2016). Annotations were pulled from the DAVID gene

annotation tool website (Jiao et al., 2012). FUNC-E was run with p-value cutoff of 0.01 (--ecut

0.01) and a high stringency (--preset high). The False Discovery Rate (FDR) adjusted Fisher’s p-

value (Fig. 2) for significantly enriched terms for the WC_KC comparison were mapped as

heatmaps using Graphpad Prism 8 (Graphpad Software Inc). FDR-adjusted Fisher’s p-values from

the WS_KS comparison were mapped to the enriched terms from the WC_KC comparison, with

non-significant annotations plotted in gray. Heatmaps of gene expression were created in

Graphpad using log2 fold change values from comparisons with both independent kea1kea2 lines

for significantly differentially expressed genes from the WC_KC consensus list (WC_KC1,

WC_KC2, FDR-adjusted p-value < 0.05) and corresponding log2 fold changes for the NaCl

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treatment comparisons (WS_KS1, WS_KS2, non-significantly expressed genes plotted in gray).

The heatmaps were then arranged in the vector graphics program Inkscape (open source).

Spectinomycin Susceptibility Assay. Plates for spectinomycin plates were prepared as described in

the plant growth section except that sterilized seeds were directly sown on treatment media.

Spectinomycin treatment plates contained 0, 1, 3, 5, and 7 µg ml-1 spectinomycin dihydrochloride.

Plants were then grown for 14 days in the light and temperature conditions described in the plant

growth section. Survival rates were calculated by dividing the number of plants with developed

true leaves by the total number of germinated seedlings for each genotype. The rap-1 genotype

was predisposed towards poor growth. Thus, the survival rate for each genotype was normalized

to the 0 µg ml-1 spectinomycin treatment. Significant differences in survival rates from the WT at

each spectinomycin concentration were determined using a Kruskal Wallace Test and Dunn’s

Multiple Comparisons test (p-value < 0.05).

Immunoblotting leaf proteins. To prepare samples, plants were grown on soil for 21-24 days as

described above. 20 mg of fresh weight from young or mature leaves was solubilized in 66 µl 4x

Laemmli buffer, then the final volume was adjusted to 200 µl with ddH2O. Samples were incubated

at 80°C for 10 minutes, then spun at 6000g. For each sample, 2.5 µl of supernatant (~ 0.25 mg

FW) was loaded onto a 12% (w/v) polyacrylamide SDS-PAGE gel and run at 100V. Proteins were

blotted from the gel onto 0.2 µm pore-size Biotrace NT nitrocellulose membrane (Pall

Corporation) and blocked for 1 hour in blocking buffer (TTBS with 5% (w/v) nonfat milk powder).

After rinsing 3 times for 5 minutes in TTBS, blots were incubated with respective primary

antibody. RbcL, RbcS, V-ATPase and Lhcb1 antibodies were acquired from Agrisera. Custom

antibodies included KEA1/2 (Bölter et al., 2019), Tic40 (Stahl et al., 1999), Tic110 (Lübeck et al.,

103

1996). Incubation was 1 hour at room temperature, or overnight at 4°C. Subsequently, blots were

rinsed in TTBS and incubated with secondary antibody (Peroxidase-conjugated Affinipure Goat

Anti-Mouse Ig (H+L), Protein Tech) diluted 1:25,000 in blocking buffer for 90 minutes. Blots

were then incubated with Clarity Western ECL Substrate (Bio-Rad) and imaged on a LICOR C-

DiGit scanner (LICOR).

In-vivo 35S labeling of plastome-encoded proteins and Pulse-Chase analysis. About 5 mg of tissue

was punched from young and mature leaves of each genotype and placed into microfuge tubes.

Punches were vacuum infiltrated with 50 µl of reaction buffer (1 mM KH2PO4 pH 6.3, 75 µM

cycloheximide) and incubated on ice in the dark for 30 minutes. Tissue was then infiltrated with

10 µCi of Express 35S Protein Labelling Mix (Perkin Elmer) and incubated at 25 °C under 300

photons m-2s-1 of white light. For the chase, samples were washed once with incubation buffer,

then once with chase buffer (reaction buffer with 10 mM L-cysteine, 10 mM L-methionine).

Samples were then vacuum infiltrated with 50 µl of chase buffer and placed back under light.

Samples were processed at each timepoint by washing twice in 200 mM Na2CO3, then

homogenized in 50 µl of 100 mM Na2CO3, mixed with 33 µl 4x Laemmli buffer, and incubated at

80°C for ten minutes. Samples where then spun at 1000g for 5 minutes. Supernatant was loaded

onto a 12% (w/v) polyacrylamide SDS-PAGE gel. Gels were then Coomassie stained, dried, and

applied to a phosphor screen for 12-16 hours. Screen was imaged on a Typhoon Phosphorimager.

The intensity of the radioactive label in different bands on the gel was quantified in ImageJ. To

quantify total labelled protein, lanes of the gel were run through a scintillation counter. Values

were normalized to fresh weight, and then to the WT control for each experiment. Means were

determined to be different from the WT control using a Holm-Sidak t-test (p-value < 0.05).

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Array-based Ribosome and Transcript Profiling of Plastome-encoded genes. Ribosome footprints

and total RNA were isolated as described in (Zoschke et al., 2013), following the modifications

introduced in (Trösch et al., 2018). RNA labelling and hybridization was performed according to

(Zoschke et al., 2013) with slight modifications: 3.5 µg purified footprints and 4 µg fragmented

total RNA from kea1kea2 and Col-0 (control and salt condition for each) were labeled with Cy3

and Cy5, respectively, using the ULS Small RNA Labelling Kit (Kreatech Diagnostics) according

to the manufacturer´s protocol. The analysis of ribosome profiling and transcriptome data was

conducted as described in (Trösch et al., 2018).

RNA Gel Blot. 300 ng total RNA was separated on agarose-formaldehyde gels as described

previously (Beick et al., 2008). Separated RNAs were transferred to Hybond-N nylon membrane

(GE Healthcare). The target transcripts were detected by hybridization at 50°C for the short (5´and

3´) and at 65°C for the coding-region probes, respectively, with PCR products, which were body-

labeled with [alpha-P32]-dCTP using random primers or sequence-specific reverse primers. In

order to prevent saturation effects for the detection of the highly abundant mature ribosomal RNA,

10 times cold probe (non-radioactive probe) was added to the hybridization reaction. The signals

were detected by exposing phosphorimager screens to the radiolabeled membranes. Signals were

quantified with Image Lab software (Bio-Rad).

In vitro Protein Translation. Reading frames for FNR and PDHe1α (At1g01090) were cloned into

pGEM®-5Zf(+) Vector. In vitro translation of protein import substrates was performed applying

the SP6 TNT coupled transcription/translation kit (Promega, Wisconsin, USA) in the presence of

35S labeled Met/Cys for 60 min at 30°C.

105

Chloroplast Isolation and Import. Seeds from the respective lines were surface sterilized and sown

on ½ MS with 1% (w/v) sucrose plates. After three weeks at 21°C, 16 h light: 8 h dark, plants were

harvested using a razor blade from one (WT) or two (kea1-1kea2-1, kea1-2kea2-2) plates

respectively. Leaf material was homogenized with a polytron mixer for 1-2 sec in 25 ml of isolation

buffer (0.3 M sorbitol, 5 mM MgCl2, 5 mM EDTA, 20 mM HEPES, 10 mM NaHCO3, pH 8.0 with

KOH). Homogenate was then filtered through 4 layers of cheesecloth and 1 layer of miracloth.

Filtrate was centrifuged for 4 min at 1500g and 4ºC. Supernatant was discarded, and pellet was

gently resuspended in 1 ml of isolation buffer and layered over a Percoll step gradient (30%, 82%

Percoll in 0.3 M sorbitol 5 mM MgCl2, 5 mM EDTA, 20 mM HEPES-NaOH pH 8.0) and

centrifuged at 2000g for 6 min. The lower band containing intact chloroplasts was carefully

removed. Intact chloroplasts were mixed in 3 volumes of wash buffer (50 mM HEPES, 0.3 M

sorbitol, 3 mM MgSO4, pH 8.0 with KOH) and centrifuged for 4 min at 1500g and 4ºC. The final

pellets were resuspended in a small volume of wash buffer and chlorophyll concentration was

determined according as previously described (Arnon, 1949). Import was conducted in wash buffer

containing 3 mM Na-ATP, 10 mM L-Met, 10 mM L-Cys, 50 mM ascorbic acid, 20 mM K-

gluconate, 10 mM NaHCO3, 0.2% (w/v) BSA plus 2 µl translation product for 8 min at 25°C.

Afterwards, chloroplasts were pelleted at 1500g, washed once in 200 µl wash buffer and then

resuspended in SDS loading buffer. Proteins were separated on SDS gels, which were Coomassie

stained, then vacuum dried and exposed on a Phosphorimager Screen for 14 h. Screens were

analyzed by a Typhoon Phosphorimager (GE Healthcare) and radioactive bands were quantified

with Image Quant (GE Healthcare). In general, WT chloroplasts equivalent to 10-15 µg

chlorophyll were used per import. Since kea1kea2 mutants have reduced chlorophyll content,

equal plastid amounts per assay were quantified based on Coomassie stained bands from the SDS-

106

PAGE using Image Quant (GE Healthcare). Bands corresponding to RbCl and the LHC proteins

were excluded from the quantification since kea1kea2 has a preexisting deficiency in these

typically high abundance proteins. For statistical evaluation, the amount of WT chloroplasts was

set to 100%, as was the imported protein. All experiments were performed from three different

chloroplast isolations.

ACKNOWLEDGEMENTS

Thanks to Drs. Ricarda Höhner, Reimo Zoschke, Mehrdad Barahimipour, Serena Schwenkert, and

Bettina Bölter for collaborative experiments in this chapter, including ionomics, plastome

microarray, northern blots, and plastid import assays. An additional thanks to Drs. John Browse,

Jim Wallis and Serena Schwenkert for access and assistance with resources for pulse-chase

analysis. Thanks to Dr. Meng Chen (UC Riverside) for providing the var2-5 point mutant line.

Additionally, we thank Dr. Juergen Soll (Ludwig Maximilian University, Munich) for providing

custom antibodies and training in protein work.

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SUPPLEMENTAL FIGURES

Supp. Fig. 3.1: Low concentrations of MgCl2 rescued kea1kea2 photosynthetic phenotype. (A) Phenotype of WT and kea1kea2 lines grown without (control), with 7.5 mM MgCl2, 25 mM MgCl2 or 75 mM NaCl. Plants grown on 75 mM MgCl2 died shortly after germination and are not shown. Lower panels display false-color image of maximum quantum yield of PSII (Fv/Fm). (B-E) Bar graph of mean photosynthetic parameter in WT and kea1kea2 under control (white bars), MgCl2 treatment (light and dark grey bars), or NaCl (white striped bars,± SEM, n = 12-15). Parameters include (B) theoretical maximum yield of PSII, Fv/Fm (C) flux through PSII, ΦII (D) flux through regulated Non-photochemical quenching, ΦNPQ, and (E) flux through non-regulated, non-photochemical quenching, ΦNO (Kramer et al., 2004; Klughammer and Schreiber, 2008) . Asterisks denote statistical significance from control value for respective phenotype. (p-value < 0.05).

108

Supp. Table 3.1: RNA sequencing statistics

genotype

NaCl Conc. (mM) Total Reads

% Aligned Reads Read 1

% Aligned Reads Read 2

Bases (GB) Read 1

Bases (GB) Read 2

% Q30 Bases Read 1

% Q30 Bases Read 2

Fold Coverage (Coding Regions)

WT 0 29,392,044 93.44 97.13 2.91 2.91 95.88 95.19 11,799x

WT 0 40,740,612 93.87 97.35 3.96 3.95 95.20 95.19 18,614x

WT 0 40,453,913 93.92 97.3 3.99 3.99 95.24 95.49 18,568x

kea1-1kea2-1

0 38,587,392 94.37 97.11 3.81 3.81 95.28 94.75 17,096x

kea1-1kea2-1

0 37,029,081 93.75 97.22 3.66 3.66 95.63 95.36 16,786x

kea1-1kea2-1

0 33,893,788 94 97.12 3.34 3.34 95.51 94.91 15,906x

kea1-2kea2-2

0 42,599,733 93.52 97.34 4.21 4.21 95.25 95.49 19,220x

kea1-2kea2-2

0 45,671,641 94.24 97.3 4.51 4.51 95.34 95.12 21,727x

kea1-2kea2-2

0 32,801,920 93.61 97.43 3.24 3.24 95.68 95.37 14,841x

WT 67.5 37,917,850 93.84 97.16 3.75 3.75 95.47 94.67 17,715x

WT 67.5 40,669,032 93.9 97.49 4.01 4.01 95.51 95.76 19,029x

WT 67.5 40,274,442 94.15 97.16 3.97 3.97 95.47 95.21 17,365x

kea1-1kea2-1

67.5 48,370,854 93.76 97.46 4.78 4.78 95.82 95.40 22,087x

kea1-1kea2-1

67.5 29,264,903 93.8 97.27 2.89 2.89 95.16 94.81 13,603x

kea1-1kea2-1

67.5 42,574,356 93.85 97.32 4.2 4.20 95.71 95.71 19,139x

kea1-2kea2-2

67.5 35,494,226 94.34 97.22 3.51 3.51 95.73 95.16 15,895x

kea1-2kea2-2

67.5 45,876,288 93.72 97.26 4.54 4.54 95.81 95.37 20,419x

kea1-2kea2-2

67.5 38,559,588 93.72 96.9 3.81 3.81 95.28 94.75 14,905x

109

Supp. Fig. 3.2: Density plot of Fragments per Kilobase of Mapped reads (FPKM) for all replicates. Plots showing log2 fold-change FPKM (x-axis) and distribution density (y-axis) for all samples sequenced. WC = WT control, KC1 = kea1-1kea2-1 control, KC2= kea1-2kea2-2 control, WS = WT under salt treatment, KS1 = kea1-1kea2-1 under salt treatment, KS2 = kea1-2kea2-2 under salt treatment.

110

Supp. Fig. 3.3: Volcano Plots of RNA Sequencing Comparisons. Plots showing log2 fold-change expression (x-axis) and -Log10 FDR-adjusted p-value (i.e., q-value, y axis) for significantly differentially expressed genes for each comparison (q-value < 0.05). Blue dots represent downregulated genes, and red dots represent upregulated genes. There was no thresholding of log2 fold-change expression.

111

Supp. Fig. 3.4: Venn Diagram of Overlapping DEGs for control and NaCl comparisons. Numbers of up (red) or down (blue) regulated DEGs that are unique or shared between different comparisons as described in Table 3.2.

112

Supp. Fig. 3.5: Replicate immunoblots. Immunoblots for steady-state levels of plastid and nuclear-encoded proteins in young (Y) vs. mature (M) leaf tissue from WT and kea1kea2 as shown in Fig. 3.12.

113

Supp. Fig. 3.6: Additional autoradiographs from the pulse-chase analyses. Leaf discs from young or mature WT and kea1kea2 leaves were harvested after 15 and 40 minutes during the pulse portion of the experiment (red labeled lanes). After chase with non-radioactive amino acids, samples were taken every 2 hours for a total of 400 minutes (black labeled lanes). Bands corresponding to rubisco large subunit (RbcL) and the D1 reaction center of PSII (PsbA) are clearly visible.

114

Supp. Fig. 3.7: Pulse-Chase in ambient light. (A) Autoradiographs of pulse-chase analysis conducted at ambient (10 µE) or moderate (250 µE) light. Leaf discs from young or mature WT and kea1kea2 leaves were harvested after 40 minutes during the pulse portion of the experiment (red labeled lanes). After chase with non-radioactive amino acids, samples were taken every 2 hours for a total of 400 minutes (black labeled lanes). Bands corresponding to rubisco large subunit (RbcL) and the D1 reaction center of PSII (PsbA) are clearly visible. (B) Quantification of total protein produced in pulse-chase experiment (top), RbcL production (middle) and PsbA (bottom). Red line represents transition from pulse to chase stage. Total protein production was measured in a scintillation counter to quantify counts per minute (CPM). For RbcL and PsbA, intensity of autoradiograph was quantified in ImageJ and normalized to the intensity of the WT at the 40-minute timepoint (± SEM, n = 3).

115

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CHAPTER 4: DISRUPTION OF PLASTID GENE EXPRESSION DUE TO LOSS OF KEA1/2 RESULTS IN ALTERED EXPRESSION OF PHOTOSYNTHESIS-ASSOCIATED NUCLEAR-ENCODED GENES (PHANGS) VIA GENOMES

UNCOUPLED 1 (GUN1) MEDIATED RETROGRADE SIGNALING

Some text, data, and figures from this chapter were submitted for publication in The Plant Cell with coauthors Rouhollah Barahimipour, Nikolay Manavski, Ricarda Höhner, Serena Schwenkert, Bettina Bölter, Takehito Inaba, Jörg Meurer, Reimo Zoschke and Hans-Henning Kunz. ABSTRACT

The plastid is a highly sensitive organelle, whose function and metabolism are easily

perturbed by abiotic stress. The division of genes encoding chloroplast proteins between the

nuclear genome and the plastome has necessitated the development of a retrograde signaling

pathway from the plastid to the nucleus to moderate gene expression in response to plastid

damage or stress. The major pathway for retrograde expression in developing chloroplasts

is mediated by the protein Genomes Uncoupled 1 (GUN1). GUN1 responds to disruption of

plastid gene expression, reactive oxygen species (ROS) and other stresses by downregulating

the expression of Photosynthesis Associated Nuclear-encoded Genes (PhANGs). The

PhANGs include many light harvesting proteins and tetrapyrrole biosynthesis enzymes

which are presumably downregulated to avoid oxidative stress. To date, no study has

investigated how loss of plastid ion transport influences plastid retrograde signaling. Thus,

the aim of this chapter is to determine if retrograde signaling is activated in response to loss

of KEA1/2. Our investigations revealed that the Golden2-like 1 (GLK1) and Golden2-like 2

(GLK2) transcription factors, which promote expression of the PhANGs, were

downregulated in kea1kea2. Subsequently, the expression of many PhANGs were also

downregulated. This is likely the result of GUN1-mediated retrograde signaling to prevent

further damage to the chloroplast. Indeed, GUN1 loss-of-function in the kea1kea2

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background has a strong negative impact on seedling growth, survival, and photosynthesis.

Yet despite obvious signs of oxidative damage, the kea1kea2gun1 mutants continue to express

GLK1 and some of the PhANGs at high levels. Taken together, this indicates the GUN1-

mediated pathway is used by the cell to respond to loss of ion homeostasis by downregulating

PhANGs and preventing greater oxidative stress.

INTRODUCTION

The chloroplast as a sensory organelle.

For years, it has been known that abiotic stresses often cause photosynthetic limitation as an initial

negative effect, thus perturbing the reduction-oxidation balance of the chloroplast (Biswal et al.,

2011). Mechanistically, this occurs due to relative over-excitation of the photosystems from excess

light, impairment of specific steps in thylakoid electron transport or impairment of carbon fixation.

Such impairments culminate in a bottle-neck effect for the dissipation of light energy captured by

the photosystems (Biswal et al., 2011; Zhu, 2016). Excess energy then induces the production of

Reactive Oxygen Species (ROS), and other stress-related metabolites in the chloroplast (Asada,

2006; Kleine and Leister, 2016). Many of these metabolites act as signaling molecules to the

remainder of the cell, thus making the chloroplast a first responder and cellular alarm system for

abiotic stress (Zhu, 2016). Stress or damage to the chloroplast induces a signaling cascade to the

nucleus to adapt gene expression, a process referred to as retrograde signaling (Kleine and Leister,

2016; Leister et al., 2017). While the exact nature of the signal(s) sent back to the nucleus remains

poorly understood, retrograde signaling has been shown to alter nuclear gene expression to prevent

oxidative damage (Cheng et al., 2011; Woodson, 2016; D'Alessandro et al., 2018; Kacprzak et al.,

2019). This process is essential for whole-cell responses, but also for chloroplast adaptation to

stress, as most chloroplast proteins are encoded in the nucleus. Abiotic stresses which are sensed

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by the chloroplast and trigger retrograde signaling include high light (Szechynska-Hebda and

Karpinski, 2013; Estavillo et al., 2011), cold (Ensminger et al., 2006), drought (Reddy et al., 2004),

exogenous salt (Miller et al., 2010), and heat stress (Sun and Guo, 2016). Additionally, treatment

of plants with plastid translation inhibitors has revealed that the plastid not only senses and

responds to photo-inhibitory stresses, but also initiates signaling in response to impairment of

Plastid Gene Expression (PGE; Koussevitzky et al., 2007). Thus, it is of particular interest to

investigate how retrograde signaling is affected by loss of chloroplast envelope antiporters KEA1

and KEA2. As shown in chapter 3, the kea1kea2 loss-of-function mutant exhibits significant

impairment of PGE, one of the triggers for retrograde signaling (Fig. 3.9-16). Furthermore, the

decreased Fv/Fm (Fig. 3.3 A-B) and altered partitioning of harvested photochemical quenching

(Fig. 3.3 C-E) indicate increased photoinhibition and decreased photosynthetic efficiency in

kea1kea2. These alterations may be a symptom of increased ROS production in the kea1kea2

mutant. Thus, loss of ion transport in kea1kea2 lines may trigger retrograde signaling via

disruption of PGE and/or ROS production. The aim of this chapter is to examine retrograde

signaling pathways in kea1kea2 to understand how the chloroplast senses and copes with

internal ion imbalance. Based on a thorough literature review, we focused our research on

elucidating the involvement of the Genomes Uncoupled 1 (GUN1) mediated signaling pathway.

Genomes Uncoupled 1: The convergence of plastid retrograde signals for biogenic control of

Nuclear Gene Expression

Over the last few decades, several independent plastid to nucleus retrograde signaling pathways

have been characterized (Mielecki et al., 2020). The majority of known pathways, including the

high light/drought-induced Sal1-PAP mediated pathway (Estavillo et al., 2011), and the

EXECUTOR1/EXECUTOR2 (EX1/2) mediated pathway (Lee et al., 2007) primarily function in

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‘operational’ control of nuclear gene expression in response to stress metabolites produced by

mature chloroplasts (Mielecki et al., 2020). Nuclear genes triggered by Sal-PAP and EX1/2 are

mostly related to reactive oxygen species scavenging and programmed cell death. Conversely,

mutant screens with various inhibitors of plastid processes revealed several genes involved in

‘biogenic’ control of nuclear gene expression by suppressing the Photosynthesis- Associated

Nuclear-Encoded Genes (PhANGs). Originally, the representative PhANG transcript assayed in

genetic screens was light harvesting complex Lhcb1 (Mochizuki et al., 2001). Mutants which had

high expression of Lhcb1 in spite of treatment with nuraflorazon (NF), an antibiotic which induces

oxidative stress by inhibiting the production photoprotective carotenoids, or lincomycin (LIN) a

PGE inhibitor were considered Genomes Uncoupled (gun) mutants (Susek et al., 1993;

Koussevitzky et al., 2007). Five of the six original gun lines discovered in these screens where

insensitive to NF, but not to LIN (Susek et al., 1993). These mutants, designated gun2 to gun6, all

had mutations in chloroplast-targeted enzymes related to tetrapyrrole biosynthesis (Susek et al.,

1993; Mochizuki et al., 2001; Woodson et al., 2011). This indicates a potential role for tetrapyrrole

intermediates in retrograde signaling, a topic which is still under debate in the community (Terry

and Bampton, 2019). The remaining gun line was unresponsive to both NF and LIN, indicating

the mutated gene was involved in mediating a response to both photogenic ROS and disruption of

PGE (Susek et al., 1993; Koussevitzky et al., 2007). This mutant was designated gun1 and had a

genetic defect in a locus encoding a stromal pentatricopeptide repeat (PPR) protein whose exact

function is still the focus of research and controversy (Pesaresi and Kim, 2019). We chose to focus

on GUN1-mediated signaling in kea1kea2 because it is the only retrograde signaling pathway

documented to respond to disruption of PGE.

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One primary mechanism by which GUN1 protects the developing plastid from stress is to suppress

the expression of the nuclear transcription factor Golden2-Like 1 (GLK1; Kakizaki et al., 2009;

Martin et al., 2016). GLK1 and its functionally redundant relative Golden2-like 2 (GLK2) are

responsible for activating transcription of the PhANGs and inducing plastid greening and

development (Waters et al., 2009). Phytochrome-mediated red/ far red light signaling (i.e., the

anterograde signaling pathway) promotes the expression of GLK1, which in turn induces the

expression of the PhANGs, driving photomorphogenesis and de-etiolation (Oh and Montgomery,

2014). The current working model is that GUN1 suppresses photomorphogenesis by

downregulating GLK1 expression when the etioplast experiences perturbation of PGE or oxidative

Fig 4.1: A model from Martin et. al. 2016 showing opposing roles of informational light and GUN1-mediated retrograde signaling in controlling expression of the PhANGs.

From Martin et. al. 2016

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stress to balance the stoichiometry of nuclear- and plastome-encoded proteins related to

photosynthesis. An excess of components related to light harvesting and chlorophyll biosynthesis

could result in further accumulation of harmful ROS and/or tetrapyrrole intermediates. Thus, the

phytochrome-mediated anterograde pathway and the GUN1-mediated retrograde pathway act in

opposition to each other in developing plastids (Fig 4.1).

Further support for this model comes from the observation that loss of GUN1 has an

additive harmful effect on plant fitness when coupled with loss-of-function of genes involved in

PGE (reviewed in Tadini et al., 2020a). Mutants with disruption of genes related to plastid

transcription, transcript editing/processing (including PPRs), plastid translation, and plastid

protein import/folding exhibit a severe reduction in photosynthetic performance and seedling death

when introgressed into the gun1 background (Tadini et al., 2020a). Not surprisingly, these mutants

exhibit the virescent phenotype which is a hallmark of PGE mutants as described in Chapter 3. As

kea1kea2 is a virescent mutant with documented delays in plastid development and gene

expression, we hypothesize that the GUN1 mediated signaling pathway is involved in

responding to loss of KEA1/2. We predict that by downregulating the GLK1 transcription factor

and its target PhANGs, GUN1 functions to prevent ROS accumulation and oxidative damage due

to imbalanced PGE in the kea1kea2 plastid. The side effect of suppression of PhANGs is that

plastids in young leaves of kea1kea2 are slow to undergo biogenesis and greening. This would

explain the virescent phenotype and delayed chloroplast development exhibited by this mutant.

GUN1: Still an enigma

It should be noted that the exact mechanisms by which GUN1 senses stress, and the nature

of the secondary messenger which carries the signal from the plastid to the nucleus is unknown.

GUN1 is a pentatricopeptide repeat (PPR) protein targeted to the chloroplast stroma, present in

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high abundance in developing leaves, and low abundance in mature leaves (Wu et al., 2018). As

described in Chapter 3, PPR proteins are organelle-targeted nuclear-encoded proteins which bind

organellar RNA to mediate RNA processing (reviewed in Barkan and Small, 2014; Manavski et

al., 2018). However, GUN1 surprisingly does not appear to have any RNA binding activity (Tadini

et al., 2016). Rather, Co-IP and yeast 2-hybrid assays have suggested that GUN1 interacts with

numerous proteins involved in PGE and plastid proteostasis. Some of these potential interactors

include the nuclear-encoded plastid polymerase (NEP) and proteins involved in plastid

transcription (Tadini et al., 2016; Tadini et al., 2020b), proteins involved in plastid RNA

processing (Zhao et al., 2019), plastid ribosomal proteins (Tadini et al., 2016), subunits of the

plastid protein import complex, and plastid chaperones (Colombo et al., 2016). Additionally, it

has been shown that GUN1 can bind heme molecules and interact with enzymes of the tetrapyrrole

pathway (Shimizu et al., 2019). Whether GUN1 binds these proteins as part of a mechanism to

sense stress and mediate retrograde signaling, or if it has a direct role in regulating the functions

of these proteins is still unclear. What is known is that the abundance of GUN1 is regulated at the

level of protein stability (Wu et al., 2018). Stresses which induce retrograde signaling suppress

Clp protease-mediated degradation of GUN1, leading to increased levels of GUN1 in the stroma.

How the accumulation of GUN1 triggers further signaling remains mysterious (Pesaresi and Kim,

2019; Mielecki et al., 2020). In summation, the elucidation of each step in the GUN1-mediated

retrograde signaling pathway is far from complete and is outside the bounds of this dissertation.

Our aim to use the well-supported information about GUN1 present in the literature to

determine if the GUN1 protein plays an important role in sensing and responding to loss of

chloroplast ion transporters.

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RESULTS

The GLK1/2 nexus is altered in response to loss of KEA1/2

As a starting point for this chapter we re-investigated our RNA-SEQ data to see if target genes

previously published to be downstream of biogenic retrograde signaling are altered in transcription

(Mielecki et al., 2020). As discussed in Chapter 3, the transcriptomics data revealed that GLK1/2

Fig. 4.2: GLK1/2 transcription factors play an important signaling role in kea1kea2. (A) Log2 fold-change values of GLK1/2 and downstream target genes when comparing the WT to two independent kea1kea2 lines under control conditions (WC_KC1 and WC_KC2 for comparisons with kea1-1kea2-1 and kea1-2kea2-2 respectively) and salt treatment (WS_KS1 and WS_KS2 for comparisons with kea1-1kea2-1 and kea1-2kea2-2 respectively). Red boxes denote statistically significant upregulation; blue boxes denote statistically significant downregulation (FDR-adjusted p-value < 0.05). Grey boxes denote gene is not significantly differentially expressed (NS, FDR adjusted p-value ≥ 0.05).

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TFs responsible for chloroplast development and PhANG expression are downregulated in

kea1kea2 under control conditions (Fig. 3.7A, GO:0009658 chloroplast organization). The

GLK1/2 downstream targets have been characterized over the last years to contain numerous

PhANGs, particularly LHC components and chlorophyll biosynthesis enzymes (Fitter et al., 2002;

Waters et al., 2009; Berry et al., 2013). Therefore, we used the gene information to draw a

quantitative GLK1/2 nexus map by plotting RNA-SEQ log2 fold change values from our WC_KC

(WT control versus kea1kea2 control) and WS_KS comparisons (salt-treated WT versus salt-

treated kea1kea2, Fig. 4.2). Indeed, we found that GLK1/2 and many of their downstream targets

were significantly downregulated in the kea1kea2 mutant under control conditions but were no

longer significant upon salt treatment (Fig 4.2). This suggests PhANG suppression via reduction

of GLK1/2 transcripts in kea1kea2 is a response to disturbed plastid ion transport and contributes

to the low content of chlorophyll and photosynthetic rates found in the mutant. Suppression of the

PhANGs is likely a coping mechanism to prevent further damage to the kea1kea2 plastids.

Altered phytochrome signaling is not the cause of GLK1/2 downregulation in kea1kea2

Phytochrome-mediated anterograde signaling is known to trigger the expression of GLK1

and GLK2 (Oh and Montgomery, 2014), while GUN-mediated retrograde signaling is known to

repress the expression of GLK1 (Kakizaki et al., 2009; Martin et al., 2016). Thus, the decreased

accumulation of GLK transcripts and downstream PhANGs in kea1kea2 could occur due to

activation of GUN1 signaling or impairment of informational light signaling. Thus, we

investigated both pathways in the kea1kea2 background to uncover which pathway is involved in

the response to loss of plastid ion transport. Based on RNA-SEQ data, neither the R/FR sensing

phytochromes PHYA/PHYB nor the retrograde signaling component GUN1 exhibit dramatic

alteration of transcript levels (Fig. 4.2). However, the FUNC-E analysis presented in Chapter 3

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showed that downstream components of red and far-red light perception (GO:0010017, Fig. 3.8B)

were enriched in the RNA-SEQ dataset. Thus, we designed experiments to test the role of these

signaling pathways in kea1kea2 directly.

To test red and far-red light signaling pathways, we setup germination experiments for the

kea1kea2 mutant under monochromatic red and far-red light and determined the hypocotyl length

as a proxy for red and far-red light signal perception. As positive controls, we included previously

characterized phytochrome-deficient mutants phyb in red light or phyA for far-red light

experiments respectively (Nagatani et al., 1993; Reed et al., 1993; Neff and Chory, 1998). We

found that the mean kea1kea2 hypocotyl length under both monochromatic light treatments was

roughly the same as in WT when normalized to a dark-grown control (Fig 4.3). Conversely, phyb

and phyA mutants failed to respond to red and far-red light respectively and revealed the

characteristically extended hypocotyl under the given light condition, proving that the

experimental conditions were correct. In summary, we did not find evidence that either red or far-

Fig. 4.3: Hypocotyl Elongation under monochromatic light for WT, kea1kea2, and phya or phyb controls. (A-B) Hypocotyl length of 5-day-old seedlings under different light conditions normalized as a percent of genotype-specific dark control. (A) Relative hypocotyl length of WT, kea11kea2, and phyb grown under white light, dark, or red light (± SEM, n = 53-83). (B) Mean relative hypocotyl length of WT, kea1kea2, and phya grown under white light, dark, or far-red light (± SEM, n = 32-81). Differing letters above the bars denote significantly different means (p < 0.05).

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red light perception is compromised in kea1kea2 double mutants. It follows that perturbed light

signaling is unlikely a factor in the observed GLK1/2 suppression.

GUN1-mediated retrograde signaling is involved in response to loss of KEA1/2.

Disruption of PGE, changes in tetrapyrrole biosynthesis, and impairment of plastid electron

transport/generation of Reactive Oxygen Species (ROS) result in downregulation of the PhANGs

via GUN1-mediated GLK1 suppression (reviewed in (Hernandez-Verdeja and Strand, 2018). In

addition to impairment of PGE (chapter 3), we also documented significantly higher H2O2

production in kea1kea2 leaves compared to WT under control conditions, suggesting that

perturbation of the light reactions in response to loss of KEA1/2 is causing ROS production at the

thylakoid membrane (Fig. 4.4).

Thus, at least two potential retrograde signaling triggers are present in the kea1kea2

chloroplast. To genetically test the role of the GUN1-mediated retrograde signaling pathway in

governing GLK1 gene expression in response to disturbed plastid ion transport, we introgressed

Fig. 4.4: DAB stain for H2O2 production shows kea1kea2 accumulates more ROS than the WT under control conditions, but not under salt treatment. (A) An RGB image of 24-day-old living plant, false color Fv/Fm image, and RGB image of DAB-stained rosette for WT and two independent kea1kea2 lines under control and salt treatment. (B) A plot of stained leaf area for all three genotypes and both treatments. Leaf area was calculated in ImageJ (± SEM, n = 14-15 ). Differing letters above the bars denote significantly different means (p-value < 0.05).

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two independent gun1 T-DNA insertion lines gun1-101 (Ruckle et al., 2007), and gun1-201

(Martín et al., 2016) into both kea1kea2 lines, and isolated gun1kea1kea2 triple mutants (Fig. 4.5).

When we germinated all genotypes side-by-side and documented their photosynthetic

capacity, we noticed a strongly aggravated phenotype in both gun1kea1kea2 lines compared to

parental mutant lines (Fig. 4.5A-D). Many gun1kea1kea2 triple mutant seedlings either died

immediately after germination or showed significantly lower Fv/Fm than kea1kea2 double or gun1

Fig. 4.5: Retrograde signaling via GENOMES UNCOUPLED 1 (GUN1) is triggered in response to loss of KEA1/KEA2 to downregulate Photosynthesis-Associated Nuclear-encoded Genes (PhANGs). (A) Schematic showing genotype position on the plate. (B) RGB and (C) false color image of maximum quantum yield of PSII (Fv/Fm) of 1-week-old plants including WT, two independent gun1 loss-of-function mutants, the two independent kea1kea2 mutants, and gun1kea1kea2 triple mutants. (D) Mean Fv/Fm for one-week old seedlings in Fig. 4.5C (± SEM, n = 4). Differing letters above the bars denote significantly different means (p-value < 0.05). (E) 3-week-old plants including WT, kea1-1kea2-1, gun1-201, and gun1-201kea1-1kea2-1 mutant. (F) Total chlorophyll content of 3-week-old plants from Fig.4.5E (± SEM, n = 12). Differing letters above the bars denote significantly different means (p-value < 0.05).

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single mutants. Interestingly, a few gun1kea1kea2 triple mutant individuals did survive. These

slow-growing individuals were visibly greener and indeed had increased total chlorophyll

compared to kea1kea2 (Fig. 4.5F, Suppl. Fig.4.1). However, mature gun1kea1kea2 individuals

did not exhibit any recovery of Fv/Fm, indicating these plants may be suffering from photodamage

or impairment of PsbA synthesis (Suppl. Fig. 4.2).

To establish that the role of GUN1 in response to loss of KEA1/2 is to suppress expression

of GLK1 and the PhANGs, we ran RT-qPCR assays on gun1kea1kea2 and its parental lines. We

chose a selection of previously published PhANGs to assay (Ruckle et al., 2007; Woodson et al.,

2013; Kacprzak et al., 2019). Our targets included Light Harvesting Complex II proteins

(LHCB1.2, LHCB2.2, LHCB2.4) tetrapyrrole biosynthesis pathway enzymes (GUN4, HEMA1),

and membrane soluble electron carrier plastocyanin (PETE). Expression of all these of genes were

also shown to be induced by GLK1/2 (Waters et al., 2009; Leister and Kleine, 2016).

Our RT-qPCR data show that GLK1 mRNA levels were suppressed in kea1kea2 double

mutants compared to the WT but recovered in gun1kea1kea2 triple mutants (Fig. 4.6A).

Furthermore, we found that some representative PhANG members downstream of GLK1, namely

LHCB1.2 and LHCB2.2 showed partially recovered gene expression, i.e., were closer to WT

mRNA levels (Fig. 4.6B-C). Two other PhANGs including light harvesting complex II protein

LHCB2.4 and PETE also showed a trend towards decreased expression in kea1kea2 and partial

recovery in gun1kea1kea2, although none of the mean values were statistically significantly

different (Fig. 4.6D, G). However, other typical PhANG members HEMA1 and GUN4 had

increased expression in kea1kea2 and gun1kea1kea2 compared to the WT (Fig. 4.6E-F). This

contradicted the trends from the RNA-SEQ experiment (Fig. 4.2), where the two genes

significantly decrease in expression compared to the WT. Thus, while a linear connection can be

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made between GUN1-mediated repression of GLK1 and LHC expression in response to loss of

KEA1/2, other PhANGs did not exhibit reliable GUN1-mediated suppression in the kea1kea2

background.

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Fig. 4.6: Transcript level expression of GLK1 and a selection of downstream PhANGs in 24-day-old WT, kea1-1kea2-1, gun1-201kea1-1kea2-1, and gun1-201. Bars represent mean percent transcript expression for given gene normalized to mean WT expression level (± SEM, n = 9 except for D and G, where n = 3). Genes assayed for expression level include (A) GLK1, (B) LHCB1.2, (C) LHCB2.2, (D) LHCB2.4, (E) GUN4, (F) HEMA1, and (G) PETE. Differing letters above the bars denote significantly different means (p-value < 0.05). For more information about calculations and normalization, please see Methods.

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DISCUSSION

Overall, these experiments indicate the importance of retrograde signaling protein GUN1 for plant

survival after loss of KEA1/2 transporters, though the direct link to nuclear gene expression is

tenuous. RNA-SEQ data from Chapter 3 revealed that expression of many nuclear-encoded genes

related to photosynthesis were downregulated in the kea1kea2 mutant. The expression of these

genes recovered in kea1kea2 when ion homeostasis is rebalanced by NaCl treatment. This

coincided with suppression of the virescent phenotype (Fig 3.3). In particular, transcription

factors GLK1/2 which promote the expression of many PhANGs were downregulated in kea1kea2

under control conditions, yet recovered under salt treatment (Fig. 4.2). GLK1/2 expression can be

promoted via informational light, or GLK1 specifically can be suppressed by GUN1 mediated

signaling. We ruled out perturbed light sensing in kea1kea2 as the reason for decreased expression

of GLK1/2 by using hypocotyl length assays in monochromatic light (Fig. 4.3). Thus, retrograde

signaling was most likely the primary mechanism altering GLK1 expression in kea1kea2.

Consequently, we investigated if GUN1-mediated signaling influenced stress response and

GLK1/PhANG expression in kea1kea2 by generating and characterizing two independent

gun1kea1kea2 triple loss-of-function mutant lines. The gun1kea1kea2 mutant seedlings were

extremely chlorotic with low Fv/Fm and high seedling mortality, indicating that GUN1 is important

for tolerating loss of KEA1/2 (Fig. 4.5B-D). Furthermore, the few gun1kea1kea2 triple mutants

which survived to maturity exhibited partial suppression of the virescent phenotype, i.e., increased

chlorophyll content. Yet, these mature individuals continue to experience photodamage as

indicated by low Fv/Fm (Fig. 4.5 E-F, Suppl. Fig. 4.2) . RT-qPCR analysis revealed that despite

continued photodamage in 24-day-old plants, GLK1 expression remained high, as did the

expression of some of the PhANGs (Fig. 4.6). However, the PhANG candidates HEMA1 and

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GUN4 did not behave as expected based on the literature or the RNA-SEQ data. Both genes

exhibited higher expression in kea1kea2 and gun1kea1kea2 lines compared to the WT (Fig. 4.6E-

F). This contrasted with the RNA-SEQ data, where both genes were significantly downregulated

in kea1kea2 (Fig. 4.2). Thus, the primary conclusion drawn from this chapter is that while GUN1

clearly plays an important role in response to loss of KEA1/2, the direct connection to PhANG

expression remains tenuous, and other pathways may be working in tandem with GUN1 to

moderate stress response in kea1kea2. Alternatively, GUN1 may have a moonlighting role in stress

response by directly regulating PGE and other stromal processes via protein-protein interactions.

The following paragraphs will discuss some of the unexpected results and potentially provide

explanations and alternative hypotheses.

One explanation for the odd pattern of PhANG expression in gun1kea1kea2 lines is that

the gene expression assays were carried out on whole rosettes of 24-day old plants. Previous

publications typically investigated the role of GUN1 in mediating PhANG expression in seedlings

and young plants, since GUN1 reaches highest levels in undeveloped plastids (Wu et al., 2018).

Studies of GLK1-induced gene expression are also typically conducted in seedlings with ongoing

plastid biogenesis (Waters et al., 2009; Leister and Kleine, 2016; Martin et al., 2016). Furthermore,

GLK1 is most highly expressed in young leaves (Klepikova et al., 2016). Thus, GUN1 and GLK1

may not have a significant role in controlling PhANG expression in older plants or developed

photosynthetic tissues. More uniform effects on PhANG expression might be quantifiable in young

plants, or specifically in young leaves of mature gun1kea1kea2 plants. This could also explain

why the suppression of PhANG expression was more robust in kea1kea2 plants from the

transcriptomics data than the RT-qPCR data. When kea1kea2 lines are grown on sterile ½ MS

media, they exhibit a more dramatic phenotype and slower development than soil-grown plants of

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an equivalent age. Indeed, kea1kea2 lines grown on soil begin to lose the virescent phenotype,

and have a higher ratio of green, healthy mature leaves compared to chlorotic young leaves (Fig.

4.5E). As we measured transcript levels in RNA isolated from whole rosettes, the high expression

of PhANGs related to chlorophyll biosynthesis, (i.e., HEMA1 and GUN4) might be coming from

mature, green leaves. Thus, we propose a future experiment to quantify PhANG expression in

younger gun1kea1kea2 and parental controls or restrict our experiments to young leaves from 24-

day-old plants.

Based on the extreme phenotype of gun1kea1kea2 seedlings, GUN1 is clearly important

for tolerating loss of KEA1/2. While there is some indication that GUN1 is responsible for

repressing the PhANGs, this may not be the exclusive mechanism by which GUN1 responds to

loss of these transporters. GUN1 could play a direct or indirect role in chloroplast PGE and

metabolism in addition to its established retrograde signaling function to suppress the PhANGs.

While this chapter and most of the literature focuses on the suppressive effect of GUN1 on PhANG

expression, it is possible that GUN1 retrograde signaling also promotes the expression of genes

which aid plastid response to PGE defects. In our RNA-SEQ dataset, kea1kea2 grown under

control conditions displayed widespread up-regulation of transcripts encoding plastid RNA

binding proteins and other key components for proper PGE (Fig. 3.7 D-H). GUN1 may be

responsible for promoting the expression of these transcripts in response to loss of PGE or other

plastid stresses. As further evidence, loss of GUN1 has been shown to decrease the accumulation

of plastid-encoded transcripts (Tadini et al., 2020b), alter rates of plastid RNA editing (Zhao et al.,

2019), and increase flux through the tetrapyrrole pathway (Shimizu et al., 2019). These effects

could very well be the result of GUN1 mediating the expression of nuclear-encoded proteins

related to PGE and the tetrapyrrole pathway. Alternatively, these results could also indicate that

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GUN1 directly influences PGE and tetrapyrrole biosynthesis via protein-protein interactions. As

stated in the introduction, GUN1 has been shown to bind an array of stromal proteins involved in

these processes. Regardless of whether GUN1 moonlights as a direct facilitator of PGE, or simply

promotes the expression of PGE-related proteins, either phenomenon would explain why

gun1kea1kea2 individuals are extremely stressed, but do not exhibit consistent upregulation of

PhANGs seen in other higher order gun1 mutants. In either scenario, loss of GUN1 would have

an additive effect on the PGE defect already exhibited by kea1kea2.

CONCLUSION

With the information deduced from our experiments, we present a model for how the retrograde

signaling pathway might operate in response to loss of KEA1/2 (Fig 4.7). In a healthy WT leaf

cell, PGE functions normally, and ROS production is low. Thus, GUN1 remains inactive, and no

retrograde signal is sent to the nucleus to suppress the expression of GLK1 and the PhANGs.

Hence, plastid biogenesis occurs normally in response to informational light, resulting in

Fig. 4.7: A proposed model for retrograde signaling in response to loss of KEA1/2.

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developed, photosynthetic chloroplasts. However, in the kea1kea2 mutant, loss of plastid ion

homeostasis or other pleiotropic effects result in impaired PGE and increased ROS production.

These signals act separately or in conjunction to trigger GUN1-mediated retrograde signaling to

the nucleus to suppress the expression of GLK1. Without sufficiently high GLK1 levels to promote

the expression of the PhANGs, plastid biogenesis is retarded, and young leaves exhibit chlorotic

under-developed chloroplasts. In turn, this causes pleiotropic effects on photosynthesis and plant

growth. Finally, the gun1kea1kea2 triple mutant initially experiences the same stresses as the

kea1kea2 mutant. However, without GUN1-mediated signaling, GLK1 expression remains high.

Thus, there is still WT-level expression of some of the PhANGs despite increasing plastid stress.

This exacerbates the imbalance of plastid and nuclear gene expression and allows for continued

generation of ROS. The accumulation of ROS and other effects of unchecked PhANG expression

is cytotoxic for many seedlings, resulting in high mortality rates. Thus GUN1-mediated signaling

is a key component to cell response to loss of chloroplast ion homeostasis. While GUN1 likely

does not directly sense ion status of the chloroplast, other documented effects resulting from loss

of ion homeostasis, i.e. disruption of PGE (Chapter 3) and increased ROS ( Fig. 4.4) production

likely trigger retrograde signaling.

MATERIALS AND METHODS

Genotyping and growth conditions: gun1kea1kea2 lines were confirmed to be homozygous using

PCR with one gene-specific primer and one T-DNA specific primer. For a full list of lines, and

primers see Appendix B. Seeds were sterilized in 70% (v/v) ethanol, plated on ½ concentration

Murashige and Skoog (MS) media with 0.8% (w/v) agar and stratified in the dark at 4° C for 48

hours (Murashige and Skoog, 1962). Plates were then placed in Percival Growth Chamber with

150 µmol photons m-2s-1 of light, 16 h: 8 h light: dark cycle. Growth temperatures were 22° C in

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the light and 18° C in the dark. Plants where grown for 7 days then either used for analysis or

transferred to soil/treatment plates until 24 days old.

PAM photosynthesis measurements Photosynthesis measurements were conducted as described in

previous chapters.

Chlorophyll Determination. About 20 mg of leaf tissue from 24-day-old plants was ground in

liquid nitrogen, then incubated in 80% (v/v) acetone for 1 hour on ice in the dark, vortexing

occasionally. Samples were then spun at maximum speed for 5 minutes at 4°C. The absorbance of

the supernatant at 646, 663, and 750 nm was measured in a spectrophotometer. The chlorophyll

content was calculated from the absorption at these wavelengths as described previously (Porra et

al., 1989).

RT-qPCR. Total RNA was extracted and treated with DNase was described for the RNA-SEQ

experiment. After removal of genomic DNA, a cDNA pools Were prepared using a Revertaid

FirstStrand cDNA synthesis kit with Oligo(DT)18 primers (ThermoFisher). RT-qPCR reactions

Were run using PerfeCta SYBR Green FastMix (QuantaBio), with 15 ng of template cDNA, and

300 nM final concentration of each primer per reaction. Some primers from previously published

research were used, while the remainder were designed using the tool (Arvidsson et al., 2008). For

lists of primers, see Supplemental Table 4.1. Three biological replicates of each sample, and two

technical replicates of each biological replicate were run in a Biorad CFX96 Real-Time

thermocycler. PCR parameters can be found in Supplemental Table 4.2. This experiment was run

independently three times, and a ΔCq value (Schmittgen and Livak, 2008) was calculated for each

reaction using SAND, UBC9 or TIP2 as a house-keeping gene, as all three genes show low variation

in expression under different light conditions (Czechowski et al., 2005). Technical replicates for

each biological replicate were averaged, and each biological replicate was normalized to the

146

average WT level of expression. These normalized values were then used to calculate the mean

and standard error of the mean (SEM) for each genotype. Significant differences in the normalized

mean were determined using an ordinary 1-way ANOVA and Tukey’s multiple Comparison test

for parametric data, or a Kruskal Wallace Test and Dunn’s Multiple Comparisons test for non-

parametric data (p-value < 0.05).

Hypocotyl Elongation Assays. Col-0 (WT), kea1-1kea2-1, kea1-2kea2-2, phyA-211, and phyB-9

seeds Were sterilized as described in the plant growth section, but then plated in 25 mm-deep petri

dishes. Seeds Were stratified for 2-3 days at 4° C. Plates where then placed in 50 µmol photons m-

2s-1 of light of red light (RL, λ = 627 nm, 14-16 h light: 8-10 h dark) in a Fytoscope (Photon

Systems Instruments) for 12 hours to stimulate germination. After 12 hours, the plates were either

transferred to 5 µmol photons m-2s-1 of constant far-red light (FRL, λ = ~750 nm) or remained in

RL as described above for 4 more days. Hypocotyl lengths Were measured by removing seedlings

from agar, photographing flat, then quantifying hypocotyl length using the Simple Nuerite Tracer

in Fiji image analysis software. Hypocotyl lengths Were then normalized to the mean of the dark

control of each genotype, and significant differences in the normalized mean where determined

using a Kruskal Wallace Test and Dunn’s Multiple Comparisons test (p-value < 0.05).

ROS Staining. H2O2 staining was carried out by incubating rosettes in 3,3'-Diaminobenzidine

(DAB), which forms a brown precipitate when reduced by H2O2 (Arsalan Daudi, 2012). Rosettes

from control and salt treated Col-0, kea1-1kea2-1, and kea1-2kea2-2 Were vacuum infiltrated with

1% (w/v) DAB as previously described (Arsalan Daudi, 2012). Infiltrated rosettes Were exposed

to 200 µmol photons m-2s-1 of white light for 8 hours to induce H2O2 production. As a negative

control, some DAB infiltrated rosettes Were incubated in the dark. Chlorophyll was bleached from

the plants by boiling in a 3:1:1 solution of ethanol, acetic acid, and glycerol for 15 minutes. The

147

% leaf area stained with DAB was quantified in Fiji image analysis software. Significant

differences in the normalized mean where determined using a using a Kruskal Wallace Test and

Dunn’s Multiple Comparisons test (p-value < 0.05).

ACKNOWLEDGEMENTS

Thank you Dr. Michael Neff (Washington State University) for providing phya and phyb loss-of-

function lines and advice. Thanks to both Dr. Neff and Dr. Helmut Kirchhoff (Washington State

University) for providing access to monochromatic light chambers. Thanks to Dr. Kiwamu Tanaka

and Matt Marcec (Washington State University) for allowing access to RT-qPCR equipment and

advice on methodology.

148

SUPPLEMENTAL FIGURES AND TABLES

Suppl. Fig. 4.2: Theoretical maximum quantum efficiency of PSII (Fv/Fm) in 24-day-old WT, kea1-1kea2-1, gun1-201kea1-1kea2-2, and gun1-201 seedlings. Means for each genotype ((± SEM, n = 12-13). Differing letters above the bars denote significantly different means (p-value < 0.05).

Supp. Fig. 4.1: Chlorophyll A, Chlorophyll B, and Chlorophyll A/B ratios in gun1, kea1kea2, and triple mutants. Mean chlorophyll (Chl) A and B content, and Chl A/B ratio for each genotype (± SEM, n = 12). Differing letters above the bars denote significantly different means (p-value < 0.05).

149

Supplemental Table 4.1: RT-qPCR Primers

Gene AGI number Function FWD primer REV primer Source

GLK1 AT2G20570 PhANG ttctaccgccatgcctaatccg actggcggtgctctaaatctcg designed in Quantprime

LHCB1.2 AT1G29910

PhANG ggacttgctttaccccggtg tcggtagcaagacccaatgg Woodson et. al. 2013

LHCB2.2 AT2G05070

PhANG gctttgtaaactcgtgattgtg tgccaaattcacatcaaacg Woodson et. al. 2013

LHCB2.4 AT3G27690

PhANG actcctcagagcatctggtacg

tttctggatcggctgagagacc

designed in Quantprime

HEMA1

AT1G58290

PhANG ggatgaggaaagcaatggaa

gaatccctccatgcttcaaa

designed in Quantprime

GUN4 AT3G59400

PhANG ctgccgtttcaaccacaaacgc

acgtcgaatatggtcgcggtttc

designed in Quantprime

PETE AT1G76100 PhANG tggtgttcgacgaagacgag agatcttgcttgcgtccaca Woodson et. al. 2013

SAND AT2G28390

reference aactctatgcagcatttgatccact tgattgcatatctttatcgccatc Czechowski et. al. 2005

TIP2 AT3G26520 reference tcgccgcttgtttcctccttag agagaccgaacgctggaattgg designed in Quantprime

UBC9 AT4G27960

reference tcacaatttccaaggtgctgc

tcatctgggtttggatccgt

Czechowski et. al. 2005

Supplemental Table 4.2: RT-qPCR Protocol Step Temperature Duration

1 95°C 0:30 2 95°C 0:15

3 60°C 0:30

4 Read plate

5 Repeat steps 2-4 50x

6 65°C 0.31

7 65°C 0.05

8 Read plate

9 Repeat steps 7-8 60x, increasing temp by 0.05°C

150

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CHAPTER 5: CONCLUSION

The overarching goal of this dissertation was to investigate the molecular basis of the

photosynthetic and developmental phenotype resulting from loss of plastid envelope-localized

K+/H+ antiporters KEA1/2 in A.

thaliana. This goal was broken down

into 3 aims (Fig. 5.1): 1) Quantify the

effect of KEA1/2 loss-of-function on

chloroplast ion concentrations; 2)

Determine if plastid gene expression

(PGE) is altered by loss of KEA1/2; 3)

Determine the pathways used by the

chloroplast to sense loss of ion

homeostasis and mediate changes in

nuclear gene expression.

As loss of KEA1/2 appears to have a

pleiotropic effect on the mutant

phenotype, we used the NaCl-

mediated rescue phenomenon of the

kea1kea2 virescent phenotype to separate the effect of loss of KEA1/2 on chloroplast development

from other effects.

Fig. 5.1: Research Aims

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After investigating these aims, we were able to build a comprehensive model of the

functional role of KEA1/2 transporters, and how their loss-of-function influences plastid

development and photosynthesis (Fig. 5.2). In a WT leaf cell (Fig. 5.2A), KEA1/2 are present,

maintaining stromal K+ levels and generally helping mediate chloroplast ion homeostasis. This

ensures that the buffer conditions of the stroma are optimal for nuclear-encoded chloroplast RNA

Binding Proteins (cRBPs) to interact with their plastome-encoded RNA targets, particularly

rRNAs. The result is proper processing of rRNAs and assembly of functional ribosomes. Thus,

plastid gene expression (PGE) occurs at rates appropriate to the metabolic status of the plastid.

Without significant PGE disturbances or accumulation of reactive oxygen species to act as triggers,

retrograde signaling via stromal mediator Genomes Uncoupled 1 (GUN1) remains inactive. This

leaves Golden2-Like (GLK) transcription factors in the nucleus unhindered to promote the

transcription of Photosynthesis-Associated Nuclear-Encoded Genes (PhANGs). The PhANGs

Fig. 5.2: Model depicting consequences of KEA1/2 loss-of-function for plastid ion homeostasis, Plastome Gene Expression (PGE), and retrograde signaling. Abbreviations are as follows = Plastome Gene Expression; PhANGs = Photosynthesis-Associated Nuclear-encoded Genes; cRBPs = chloroplast RNA-Binding Proteins.

157

include genes encoding light harvesting complexes and chlorophyll biosynthesis enzymes, among

others. Expression of the PhANGs governs etioplast development into mature, photosynthetic

chloroplasts.

In leaf cells of kea1kea2 mutants (Fig. 5.2B), loss of K+/H+ exchange across the envelope

results in K+ accumulation in the stroma and disruption of other ion gradients. This creates

unfavorable conditions for cRBP-RNA interactions, resulting in rRNA processing defects. These

rRNA processing defects and independent effects of stromal ion concentration on ribosomal

assembly and function result in lower translation rates, and lower overall rates of PGE. Lower

PGE, likely in combination with ROS accumulation, triggers GUN1- mediated retrograde

signaling to prevent further damage to the plastid by repressing the expression of GLK1.

Conseqeuntly, the expression of the PhANGs is decreased, resulting in delayed plastid

development and lower photosynthetic rates in kea1kea2. The macroscale effect is a virescent

phenotype in kea1kea2 mutants.

When treated with exogenous sodium chloride (NaCl), K homeostasis is partially restored

in kea1kea2 mutants (Fig. 5.2C). This is more favourable for stromal cRBP-RNA interactions, and

partially restores rRNA processing and ribosomal assembly/function. The result is sufficeint

reocovery of PGE to keep GUN1-mediated retrograde signaling inactive and allow for expression

of GLK1 and downstream PhANGs. Thus, plastid development proceeds at almost WT-levels in

NaCl-treated kea1kea2 mutants, preventing the virescent phenotype.

While further experiments to better characterize the plastid ionome, directly quantify

protein-RNA binding in the stroma, and characterize ribosomal assembly in the kea1kea2 mutant

are nessecery, our results suggest a new role for ion transporters in bioenergetic organelles. In

addition to regulating biophysical phenomena such as Proton Motive Force (PMF), and

158

transporting cofactors for enzymes and electron transport protiens, ion transporters are responsible

for maintaining the optimal conditions required for flawless organelle gene expression. These new

findings have broader impacts on how we understand chloroplast function, and therefore will be

useful in the future for any attempts at re-engineering photosynthesis. Furthermore, our research

underscores the intricate mechanims photosynthetic organisms have evolved to cope with internal

and external stresses, many of which are sensed initially by pertubation of processes within the

plastid.

On a personal note, I’d like to express my gratitude to my advisor for his time, support,

and good sense of humour. Your mentorship has changed my life for the better. I’d also like to

thank my committee members for your feedback and advice, which have allowed me to grow as a

scientist. A big thanks to all of our collaberators for sharing their time, effort and expertise to make

this project possible. Finally, I give my gratitude to my labmates, and the students, faculty, and

staff associated with the Molecular Plants Sciences program at Washington State University. I

cannot imagine how my life would be if I had not spent the past 5 years with your positive influence

and encouragement.

159

APPENDIX A: THEORETICAL OVERVIEW OF SELECTED METHODS

This dissertation features a range of analytical, ecophysiological, and biochemical techniques to

characterize the KEA1/2 loss-of -function mutant. While some methods are common-place and can

be considered general knowledge, others are specific to certain subfields of research. Thus, I will

provide a brief theoretical overview of some of the more obscure methods I use or reference to

facilitate the understanding of my results. Practical information and protocols related to these

techniques can be found in the designated METHODS section in each chapter.

Elemental analysis methods

Most elemental analysis methods that are based on atomic spectrometry- the quantification of how

atoms in a substance interact with or emit electromagnetic radiation. For optical spectrometry,

elements in a sample are identified based on their characteristic emission of specific wavelengths

of light. Mass spectrometry (MS) quantifies elements based on mass/charge ratios. Both types of

atomic spectrometry require ionization of atoms in the sample to quantify optical properties or

separate particles by mass-charge.

Fig. A1: Three common methods of elemental analysis. From top to bottom: Atomic Absorption Spectroscopy (AAS), Inductively Coupled Plasma Emission Spectroscopy (ICP-AES) and Mass Spectrometry (ICP-MS). Graphic from (Thomas, 2015).

160

Inductively Coupled Plasma (ICP)-based methods. In Inductively Coupled Plasma (ICP) based

methods, sample is nebulized into argon carrier gas, then ionized and excited from ground state by

passing through a high-temperature plasma flame. For ICP combined with Atomic Emission

Spectroscopy (ICP-AES; sometimes also referred to as “Optical Emission Spectroscopy” or ICP-

OES), elemental composition of a sample is determined based on characteristic wavelengths of

fluorescence emitted by atoms (Thomas, 2015). In ICP-MS instruments, ionized sample is injected

into a mass analyzer (Hann et al., 2015; Thomas, 2015; Wilschefski and Baxter, 2019). The

advantage of both these methods is that they can quantify a wide range of elements at one time,

usually with very low limits of detection. As described in chapter 2, the disadvantage of these

methods is that they require liquid samples for nebulization, therefore special sample preparation

is necessary for analysis of solid biological materials (Husted et al., 2011; Maathuis and Maathuis,

2013).

Flame Photometry. Flame photometry is an old method which works on a similar principle as ICP-

AES, using emission of characteristic wavelengths of fluorescence to identify and quantify

different elements (Barnes et al., 1945). However, it typically uses a flame that is not hot enough

to excite many elements, so its use is limited.

Atomic Absorption Spectroscopy (AAS). Unlike ICP-AES and flame photometry, Atomic

Absorption Spectroscopy (AAS) measures the absorption of specific wavelengths of light by

ground-state atoms, rather than emission spectra of excited atoms to quantify an element in a

sample. Each element is measured at a unique wavelength of light, provided by a lamp in the AAS

instrument. AAS is particularly useful for measuring light elements such as sodium (Na) which

are difficult to measure using emission spectra. However, as each element requires a different light

161

bulb, this technique can only measure one element at a time (Isaac and Kerber, 1971; Thomas,

2015).

X-Ray Fluorescence (XRF). X-ray fluorescence (XRF) based methods (including Total-Reflection

X-ray Fluorescence) are like ICP-AES in that they quantify element concentrations by measuring

emitted fluorescence from excited atoms. However, these methods use a high-energy X-ray beam

to excite the sample rather than a plasma torch (Bohlen and Reinhold, 2015). The original XRF

method has a major disadvantage- fluorescence from elements of interest could be absorbed or

induced by other molecules in the sample during X-ray excitation, i.e., matrix effects (Bowers,

2019). The TXRF method overcomes this limitation by applying the X-ray beam to a thin layer of

sample at a glancing angle and placing the detector directly above the sample (Fig. A2). This

allows for quantification of analytes without significant spectral noise due to the matrix effect.

Fig A2: Schematic of typical TXRF setup. Graphic from Bruker Corp.

162

Pulse-Amplitude Modulated (PAM) Chlorophyll Fluorescence

This dissertation makes ample use of Pulse-

Amplitude Modulated (PAM) chlorophyll

fluorescence to non-destructively characterize

photosynthetic efficiency in A. thaliana. This

method measures the fluorescence of

chlorophyll A (Chl A) pigment in photosystem

II antenna to determine how captured light

energy is used. The current model of

photosynthesis assumes light energy captured

by chlorophyll has three potential fates: (1)

Transfer to the D1 reaction center where it is used to split water and produce an excited electron,

i.e. photochemistry; (2) dissipation of energy through nonphotochemical quenching (NPQ); and

(3) re-emission as chlorophyll fluorescence (Maxwell and Johnson, 2000; Baker, 2008). NPQ

mechanisms can be regulated or non-regulated. Regulated NPQ is complex, but in essence is a set

of mechanisms used by the plant to safely dissipate excess energy as heat (Ruban, 2016).

Nonregulated NPQ is dissipation of energy through damage to the reaction center of PSII, i.e.

photodamage (Klughammer and Schreiber, 2008).

The proportion of total absorbed light energy used in a particular process is referred to as

a ‘quantum yield’, designated by the Greek character Φ. Measuring the quantum yield of

fluorescence (ΦF) in scenarios which favor or inhibit different means of energy dissipation allow

estimation of the quantum yields of photochemistry (ΦII), regulated NPQ (ΦNPQ), and

photodamage (ΦNO) (Kramer et al., 2004; Klughammer and Schreiber, 2008). Mechanistically,

Fig. A3: The three fates of light energy absorbed by PSII. Figure from (Baker, 2008).

163

this is accomplished using a device which includes a modulated (rapidly pulsed) light source of

low intensity to induce chlorophyll fluorescence without starting photosynthetic electron transfer.

A specially engineered camera which is tuned to measure fluorescence induced by the modulated

light. The device is also equipped with another, high intensity light source which can subject

specimens to a specific regimen of photosynthetically active (i.e., actinic) light treatment.

While the intensity and pattern of light treatments and pulses may vary, a typical regimen

begins by first estimating chlorophyll fluorescence in dark-adapted plants with the modulated light

to determine the base level of fluorescence when almost all absorbed light is directed to

photochemistry. Then, fluorescence is measured when dark-adapted plants are subjected to a burst

of intense actinic light, a scenario where fluorescence quenching by photochemistry and NPQ are

Fig. A4: An example of a chlorophyll fluorescence spectra and the meaning of various peaks. Figure from Baker 2008.

164

negligible, and all absorbed energy is partitioned to photodamage or fluorescence. After the initial

intense burst of actinic light, an actinic light of environmentally relevant intensity will switch on

for several minutes to promote the induction of NPQ and photochemistry. For the duration that

this light is on, fluorescence is measured, and intense bursts of light are applied periodically to

block photochemistry by completely reducing the plastoquinone pool. An example of a typical

PAM chlorophyll fluorescence spectra is shown in Fig. A4. The quantum yields ΦII, ΦNPQ, and

ΦNO can be calculated for different times points during the light regimen by taking ratios of

various fluorescence peaks in the spectra. It should be noted that there are many other parameters

that can be calculated using PAM chlorophyll fluorescence values (for a review of parameters and

equations, see Maxwell and Johnson, 2000; Baker, 2008).

For my purposes, I focus on steady-state values of ΦII, ΦNPQ, and ΦNO measured under

light conditions that replicate my plants’ growing conditions. High values of ΦII relative to ΦNPQ,

and ΦNO indicate a plant has high photosynthetic efficiency under given light conditions. I also

typically measure the initial ΦII in dark-adapted plants, which represents is the maximum quantum

yield of PSII (Fv/Fm). This parameter is frequently measured in ecophysiology studies, as low

values often indicate a plant suffers from irrecoverable photodamage, which is a common symptom

of plant stress (Maxwell and Johnson, 2000; Ruban, 2016).

165

WORKS CITED Baker, N.R. (2008). Chlorophyll Fluorescence: A Probe of Photosynthesis In Vivo. Annual review of plant biology 59, 89-113. Barnes, R.B., Richardson, D., Berry, J.W., and Hood, R.L. (1945). Flame Photometry A Rapid

Analytical Procedure. Industrial & Engineering Chemistry Analytical Edition 17, 605-611.

Bohlen, A.v., and Reinhold, K.m. (2015). Total-reflection x-ray fluorescence analysis and related

methods. (Hoboken, New Jersey: Hoboken, New Jersey : Wiley). Bowers, C. (2019). Matrix Effect Corrections in X-ray Fluorescence Spectrometry. Journal of

Chemical Education 96, 2597-2599. Hann, S., Dernovics, M., and Koellensperger, G. (2015). Elemental analysis in biotechnology.

Current Opinion in Biotechnology 31, 93-100. Husted, S., Persson, D.P., Laursen, K.H., Hansen, T.H., Pedas, P., Schiller, M., Hegelund, J.N.,

and Schjoerring, J.K. (2011). Review: The role of atomic spectrometry in plant science. J. Anal. At. Spectrom. 26, 52-79.

Isaac, R.A., and Kerber, J.D. (1971). Atomic Absorption and Flame Photometry: Techniques and

Uses in Soil, Plant, and Water Analysis (Madison, WI, USA: Madison, WI, USA: Soil Science Society of America), pp. 17-37.

Klughammer, C., and Schreiber, U. (2008). Complementary PS II quantum yields calculated

from simple fluorescence parameters measured by PAM fluorometry and the Saturation Pulse method PAM Appl. 1, 27-35.

Kramer, D.M., Johnson, G., Kiirats, O., and Edwards, G.E. (2004). New Fluorescence

Parameters for the Determination of QA Redox State and Excitation Energy Fluxes. Photosynth Res 79, 209.

Maathuis, F.J.M., and Maathuis, F.J.M. (2013). Plant Mineral Nutrients Methods and Protocols.

(Totowa, NJ: Totowa, NJ : Humana Press : Imprint: Humana). Maxwell, K., and Johnson, G. (2000). Chlorophyll fluorescence - a practical guide. J Exp Bot 51,

659 - 668. Ruban, A.V. (2016). Nonphotochemical Chlorophyll Fluorescence Quenching: Mechanism and

Effectiveness in Protecting Plants from Photodamage. Plant Physiol. 170, 1903-1916. Thomas, R. (2015). Determining elemental impurities in pharmaceutical materials: how to

choose the right technique. Spectroscopy (Springfield, Or.) 30, 30.

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Wilschefski, S.C., and Baxter, M.R. (2019). Inductively Coupled Plasma Mass Spectrometry:

Introduction to Analytical Aspects. Clin Biochem Rev 40, 115-133.

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APPENDIX B: MUTANT LINES AND GENOTYPING PRIMERS

Polymorphism

Type Stock FWD

primer REV

primer tDNA

primer Reference

kea1-1 T-DNA insertion

SAIL_586_D02

gcaattattgcagtaatagccactg

c

ccctcaaactcctacaatttctatg

tagcatctgaatttcataaccaatct

cgatacac

Kunz, Gierth et al. 2014

kea1-2 T-DNA insertion

SAIL_1156_H07

See above See above See above See above

kea2-1 T-DNA insertion

SALK_045234

gttgctatcactggcataattgc

gatagcgagtgtgccttcaataatc

tgg

attttgccgatttcggaac

See above

kea2-2 T-DNA insertion

SALK_009732

ggatttacacttcttggggcagg

ctaagcctttcgacagagag

attttgccgatttcggaac

See above

phya-211

γ ray CS6223 N/A

Nagatani, Reed et al. 1993

phyb-9 EMS

mutagenesis CS6217 N/A

Reed, Nagpal et al. 1993

gun1-101

T-DNA insertion

SAIL_33_D01

gtgggttctgctgtttctttg

ccaaacattgttaggaccattgg

tagcatctgaatttcataaccaatct

cgatacac

Ruckle, DeMarco et al. 2007

gun1-201

T-DNA insertion

SAIL_290_D09

gtgggttctgctgtttctttg

atgctgcatatcagatttcgg

tagcatctgaatttcataaccaatct

cgatacac

Martin, Leivar et al. 2016

rap-1 T-DNA insertion

SAIL_1223 See Kleinknecht et al. 2014

rps5 T-DNA insertion

SALK_095863

agcagatttctgaacagcagc

aattaacgttgctcgttggtg

attttgccgatttcggaac

Zhang, Yuan et al. 2016

var2-5 EMS

mutagenesis

Yu et al. 2008