1 running head: live imaging of inorganic phosphate 2 3 · 1 live imaging of inorganic phosphate in...

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Running head: Live Imaging of Inorganic Phosphate 1

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Corresponding author: 3

Wayne K. Versaw 4

Department of Biology 5

Texas A&M University 6

College Station, Texas, 77843 USA 7

Tel. 979-847-8587 8

Fax: 979-845-2891 9

E-mail [email protected] 10

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Research area: Cell Biology 12

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Plant Physiology Preview. Published on January 26, 2015, as DOI:10.1104/pp.114.254003

Copyright 2015 by the American Society of Plant Biologists

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Live Imaging of Inorganic Phosphate in Plants with Cellular and Subcellular 1

Resolution 2

Pallavi Mukherjee, Swayoma Banerjee, Amanda Wheeler, Lyndsay A. Ratliff, Sonia 3

Irigoyen, L. Rene Garcia, Steve W. Lockless, Wayne K. Versaw* 4

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Department of Biology, Texas A&M University, College Station, Texas, USA 6

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One-sentence summary: 9

Genetically encoded sensors enable dynamic monitoring of phosphate concentrations in 10

different cells and cell compartments of live plants. 11

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FOOTNOTES 1 1This work was supported by funds from the National Science Foundation (IOS-2

0956486) to W.K.V., the Welch Foundation (A-1742) to S.W.L., and the Howard 3

Hughes Medical Institute to L.R.G. 4

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*Corresponding author: [email protected] 6

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ABSTRACT 1

Despite variable and often scarce supplies of inorganic phosphate (Pi) from 2

soils, plants must distribute appropriate amounts of Pi to each cell and subcellular 3

compartment to sustain essential metabolic activities. The ability to monitor Pi dynamics 4

with subcellular resolution in live plants is therefore critical for understanding how this 5

essential nutrient is acquired, mobilized, recycled and stored. Fluorescence indicator for 6

Pi (FLIPPi) sensors are genetically encoded fluorescence resonance energy transfer 7

(FRET)-based sensors that have been used to monitor Pi dynamics in cultured animal 8

cells. Here we present a series of Pi sensors optimized for use in plants. Substitution of 9

the eYFP component of a FLIPPi sensor with a circularly permuted version of Venus, 10

cpVenus, enhanced sensor dynamic range nearly 2.5-fold. The resulting cpFLIPPi sensor 11

was subjected to a high-efficiency mutagenesis strategy that relied on statistical coupling 12

analysis to identify regions of the protein likely to influence Pi affinity. A series of 13

affinity mutants was selected with Kd values of 0.08-11 mM, which spans the range for 14

most plant cell compartments. The sensors were expressed in Arabidopsis thaliana, and 15

ratiometric imaging was used to monitor cytosolic Pi dynamics in root cells in response 16

to Pi deprivation and resupply. Moreover, plastid-targeted versions of the sensors 17

expressed in wild type and a mutant lacking the PHT4;2 plastidic Pi transporter 18

confirmed a physiological role for this transporter in Pi export from root plastids. These 19

cpFLIPPi sensors therefore enable detailed analysis of Pi dynamics with subcellular 20

resolution in live plants. 21

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23

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INTRODUCTION 1

Phosphorus is an essential element that plants acquire and assimilate in the form 2

of inorganic phosphate (Pi). This macronutrient is a component of numerous metabolites 3

and macromolecules, including ATP, nucleic acids and phospholipids, and serves key 4

roles in energy transfer reactions, signal transduction processes and the regulation of 5

enzyme activities. Of fundamental importance to plants, Pi also serves critical roles in 6

photosynthesis, both as a substrate for ATP synthesis via photophosphorylation and as a 7

regulator in the partitioning of fixed carbon between the starch and sucrose biosynthetic 8

pathways. 9

In many soils, particularly those used for low-input agriculture, the amounts of Pi 10

available to plants are limiting for growth and productivity (Vance et al., 2003). Most of 11

the Pi in soils is unavailable because it is immobilized through formation of insoluble 12

complexes or exists in organic forms such as phytate that plants cannot use directly 13

(Schachtman et al., 1998). As a result, concentrations of free Pi in soil solution range 14

from 1 to 10 μM, whereas cells require Pi in the millimolar range (Bieleski, 1973). 15

To acclimate to Pi limitation, plants have evolved mechanisms to enhance Pi 16

acquisition and also to mobilize, recycle and conserve internal stores. These mechanisms 17

include secretion of organic acids and phosphatases (Vance et al., 2003), increased 18

growth of lateral roots and root hairs (Bates and Lynch, 2000; Peret et al., 2011), 19

production of high-affinity Pi transporters at the root-soil interface (Misson et al., 2004; 20

Shin et al., 2004), formation of symbiotic association with mycorrhizal fungi, which 21

enhance Pi scavenging capabilities (Javot et al., 2007), modification of metabolic 22

pathways (Plaxton and Tran, 2011), and altered patterns of Pi translocation between 23

organs and transport between subcellular compartments (Walker and Sivak, 1986; 24

Mimura, 1999; Raghothama, 1999). Substantial insights have been gained into the 25

underlying biochemical identities and regulatory strategies for such adaptive responses, 26

including those related to sensing and signaling of Pi status (Rouached et al., 2010; 27

Chiou and Lin, 2011; Plaxton and Tran, 2011; Jain et al., 2012; Liu et al., 2014; Zhang et 28

al., 2014). However, a thorough understanding of their respective mechanisms and how 29



6

these are integrated is limited by the inability to assess intracellular Pi concentrations 1

with high spatial and temporal resolution. 2

Genetically encoded fluorescent sensors, or biosensors, have proven to be 3

powerful tools for monitoring metabolites and ions in vivo because their expression and 4

subcellular targeting can be manipulated and fluorescence imaging is nondestructive 5

(Lalonde et al., 2005; Okumoto et al., 2012). Sensor proteins are fusions of a ligand-6

binding domain or protein with one or two fluorescent proteins, e.g., GFP and related 7

variants. Sensors with a single fluorescent protein report ligand-dependent changes in 8

conformation as changes in fluorescence intensity, whereas sensors with two fluorescent 9

proteins can yield changes in Förster (or fluorescence) resonance energy transfer 10

(FRET), which can be quantified through ratiometric imaging. FRET-based sensors have 11

been used in live plants to assess a variety of analytes including glucose, maltose, 12

sucrose, glutamine, calcium, zinc and pH (Deuschle et al., 2006; Chaudhuri et al., 2008; 13

Kaper et al., 2008; Rincon-Zachary et al., 2010; Chaudhuri et al., 2011; Adams et al., 14

2012; Gjetting et al., 2012; Krebs et al., 2012; Gjetting et al., 2013). 15

Gu et al. (Gu et al., 2006) engineered FRET-based Pi sensors named 16

fluorescence indicator protein for Pi (FLIPPi) that consist of a cyanobacterial Pi binding 17

protein fused to eCFP and eYFP, and demonstrated the utility of one of these sensors for 18

monitoring cytosolic Pi in cultured animal cells. In this study, we generated a series of 19

second-generation FLIPPi sensors that were modified and optimized for use in live 20

plants. Substitution of eYFP with a circularly permuted form of the fluorescent protein 21

Venus (cpVenus) (Nagai et al., 2002; Nagai et al., 2004) greatly increased the magnitude 22

of Pi-dependent FRET responses. In keeping with the initial nomenclature, Pi sensors 23

constructed with cpVenus were designated cpFLIPPi. We also used a targeted 24

mutagenesis approach to obtain cpFLIPPi sensors with Pi binding affinities that spanned 25

the physiological range of most cell compartments and expressed these in Arabidopsis. 26

Confocal microscopy coupled with ratiometric analysis or acceptor photobleaching 27

detected changes in cytosolic Pi levels in root epidermal cells in response to Pi 28

starvation, and these changes were fully reversed by Pi replenishment. Plastid-localized 29



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versions of the same sensors expressed in wild-type plants and in mutants lacking the 1

PHT4;2 plastidic Pi transporter (Irigoyen et al., 2011) were used to confirm a role for 2

this transporter in the export of Pi from root plastids. These results demonstrate the 3

utility of cpFLIPPi sensors for monitoring Pi distributions with both cellular and 4

subcellular resolution in live plants. 5

6

RESULTS 7

Efficacy of FLIPPi Sensors in Plants 8

In an initial attempt to monitor Pi distributions in live Arabidopsis plants we 9

generated stable transgenic lines for two of the previously described Pi sensors, FLIPPi-10

200μ and FLIPPi-30m (Gu et al., 2006). When assayed in vitro, these sensors yielded 11

robust Pi-dependent FRET responses with Kd values for Pi binding of 200 μM and 30 12

mM, respectively. The FLIPPi-30m sensor was also used to report changes in cytosolic 13

Pi concentrations in cultured animal cells (Gu et al., 2006). It should be noted that these 14

sensors exhibit a Pi-dependent decrease in FRET (Fig. 1A). 15

Constitutive expression of the sensor genes was directed from the 35S promoter, 16

and because no organellar targeting sequences were incorporated in the constructs the 17

proteins were restricted to the cytosol. Independent transgenic lines for both sensors 18

exhibited weak or highly variable fluorescence that diminished rapidly with plant age, 19

suggestive of transgene silencing (Deuschle et al., 2006). Although signal intensity and 20

stability improved when the sensors were expressed in an siRNA-defective mutant host, 21

suppressor of gene silencing 3-13 (sgs3-13) (Kumakura et al., 2009), no changes in 22

FRET were detected when plants were starved for Pi. 23

We hypothesized that one or more attributes of the FLIPPi sensors limited their 24

efficacy when used in plants. For example, if the magnitudes of Pi-dependent responses 25

were small, then subtle changes in cytosolic Pi concentrations would not be detected. 26

Similarly, no change in FRET would be expected if binding affinities for Pi were too far 27

above or below the cytosolic concentrations. To explore these possibilities we first 28

confirmed that purified FLIPPi-200μ and FLIPPi-30m sensor proteins yielded Pi-29



8

dependent FRET responses equivalent to those reported previously (Gu et al., 2006). We 1

then examined FRET responses for the purified proteins in a pseudocytosol medium 2

(Messerli and Robinson, 1998; Feijo et al., 1999; Cardenas et al., 2008) to simulate in 3

vivo conditions more closely than the buffer solution used previously (Gu et al., 2006). 4

In this case, the FLIPPi-30m sensor yielded erratic FRET signals regardless of Pi 5

concentration, which may explain the absence of Pi-dependent FRET responses in the 6

corresponding transgenic plants. In contrast, FLIPPi-200μ exhibited the expected Pi-7

dependent reduction in FRET, and the 190 μM Kd was nearly identical to that measured 8

in buffer (Fig. 1B). However, the dynamic range, i.e., the maximum change in FRET 9

ratio (∆Rmax), was relatively small (Fig. 1B), and the change was very small over the 2.5 10

to 10 mM concentration range that encompasses most estimates for cytosolic Pi in plants 11

grown in Pi-replete conditions. (Rebeille et al., 1984; Stitt et al., 1988; Lee and Ratcliffe, 12

1993; Copeland and Zammit, 1994; Gout et al., 2011). Based on these results, we chose 13

to optimize FLIPPi-200μ. 14

15

Expanded Dynamic Range of a Pi sensor with a Circularly Permuted Venus 16

Substitution of the YFP acceptor component of a FRET-based sensor for Ca2+ 17

with a circularly permuted version of Venus (cpVenus) improved dynamic range and 18

stability and reduced sensitivity to acidic pH (Nagai et al., 2004). This modified sensor 19

has been effective for monitoring Ca2+ dynamics in plants (Rincon-Zachary et al., 2010; 20

Krebs et al., 2012). To test whether this modification would also increase the dynamic 21

range of a Pi sensor we replaced the eYFP portion of FLIPPi-200μ with cp173Venus, 22

the same cpVenus variant used to improve the YC3.60 Ca2+ sensor (Nagai et al., 2004). 23

As shown in Fig. 1, the modified Pi sensor, cpFLIPPi-200μ, exhibited greater FRET 24

ratios in the absence of Pi and at all concentrations tested. More importantly, the 25

dynamic range of the modified sensor (∆Rmax -1.08) was nearly 2.5-fold greater than that 26

of the original sensor (∆Rmax -0.44). The emission spectra for cpFLIPPi-200μ (with 27

eCFP excitation) indicated that the Pi-dependent response of this sensor was due to a 28

change in FRET (increase in eCFP emission coupled to a decrease in cpVenus emission) 29



9

rather than unrelated changes in the fluorescence intensities of either eCFP or cpVenus 1

alone (Suppl. Fig. 1). 2

3

Mutant cpFLIPPi Sensor Variants with Altered Affinities for Pi 4

Although the use of cpVenus increased sensor dynamic range it had no 5

significant effect on affinity for Pi. Consequently, the magnitude of the response change 6

over the target concentration range of 2.5 to 10 mM remained small for cpFLIPPi-200μ 7

(Fig. 1). This situation necessitated a mutant screen to obtain cpFLIPPi variants with 8

altered binding affinities. 9

To limit the scope of mutagenesis we used statistical coupling analysis (SCA), a 10

sequence-based method for identifying protein residues or regions responsible for ligand 11

binding affinity and potential allosteric sites (Lockless and Ranganathan, 1999; Socolich 12

et al., 2005). SCA was conducted with an alignment of the Synechococcus Pi binding 13

protein (PiBP) sequence with 1,020 of the most closely related protein sequences (PSI-14

BLAST e-score <1e-10) available from GenBank. We found that the region spanning 15

amino acids 18 to 23 contained multiple coupled sites that map near the predicted Pi 16

binding site (Suppl. Figs. 2 and 3). We predicted that changes in this region would affect 17

the Pi binding affinity of the sensor protein. 18

We used PCR to introduce random mutations in the corresponding region of the 19

cpFLIPPi-200μ plasmid clone. The mutant library was transformed into E. coli, and 20

lysates of individual clones were screened for Pi-dependent FRET. The mutagenesis 21

strategy proved to be remarkably effective. Of the nearly 600 mutants screened, less than 22

1% resembled the parent sensor and 80% of the mutants exhibited Pi-dependent FRET 23

with altered affinity. The remaining 20% of the mutant population were either 24

unresponsive to Pi or had ratiometric signals that could not be accurately fit to a binding 25

isotherm. Eight candidate sensor mutants were selected, and their Pi-dependent FRET 26

responses were verified using the purified proteins (Table I). The Kd values ranged from 27

0.08 to 11 mM, which spans the predicted physiological range for the cytosol and other 28

cell compartments (Rebeille et al., 1984; Stitt et al., 1988; Sharkey and Vanderveer, 29



10

1989; Lee and Ratcliffe, 1993; Copeland and Zammit, 1994; Gout et al., 2011; Tiessen 1

et al., 2012). 2

3

Specificity and pH Sensitivity of cpFLIPPi Sensors 4

To determine whether cpFLIPPi mutations that altered affinity for Pi also 5

affected specificity we examined the effect of other oxyanions, including 6

organophosphates, on FRET responses. None of the sensors showed responses indicative 7

of binding to sulfate or nitrate, and results for one sensor, cpFLIPPi-6.4m, are shown in 8

Supp. Fig. S4A. Similarly, treatment with glycerol-2-phosphate and pyrophosphate, a 9

phosphate ester and anhydride, respectively, yielded little or no FRET responses (Supp. 10

Fig. S4B). These results suggest that the cpFLIPPi sensors have retained high specificity 11

for Pi. 12

Our in vitro calibration assays and related tests for specificity were conducted at 13

pH 7.5 to simulate the cytosol (Pratt et al., 2009; Gout et al., 2011). Although the cytosol 14

is tightly buffered at this pH, the chloroplast stroma can vary from pH 7 to 8 in response 15

to light (Demmig and Gimmler, 1983). Because one of our goals was to monitor 16

plastidic Pi concentrations, we examined Pi-dependent FRET responses over this pH 17

range for each of the cpFLIPPi sensors. We observed a consistent trend in which affinity 18

decreased with increasing pH and results for one representative sensor, cpFLIPPi-6.4m, 19

are shown in Fig. 2. The apparent differences in affinity over this pH range can be 20

largely attributed to the effect of pH on ionization of Pi with specificity for the 21

monobasic form. Unfortunately, these sensors had very low dynamic ranges when 22

assayed at pH 5.5, which would limit their utility in acidic compartments like the 23

vacuole and apoplast. 24

25

Subcellular Targeting of cpFLIPPi Sensors in Arabidopsis 26

To test the fidelity of targeting cpFLIPPi sensors to different compartments of 27

plant cells we constructed expression clones for each of the eight different cpFLIPPi 28

sensors to yield proteins with and without an N-terminal RbcS chloroplast transit peptide 29



11

(Lee et al., 2006). To avoid potential complications associated with the 35S promoter, 1

the Arabidopsis UBQ10 promoter was used to drive constitutive expression at moderate 2

levels (Norris et al., 1993; Geldner et al., 2009; Grefen et al., 2010). Constructs were 3

introduced into the siRNA mutant sgs3-13 (Kumakura et al., 2009) to minimize potential 4

loss of fluorescent signals due to post-transcriptional gene silencing (Deuschle et al., 5

2006). Protoplasts were prepared from leaves of 5-week-old plants and fluorescent 6

signals were viewed with confocal microscopy. Co-localization with chlorophyll 7

autofluorescence shown in Fig. 3 confirmed that addition of the RbcS chloroplast transit 8

peptide was sufficient to target sensors to plastids, whereas unmodified sensors were 9

excluded from plastids and appeared to be restricted to the cytosol. FRET emission was 10

detected with the same localization patterns in palisade mesophyll cells of intact leaves 11

(Supp. Fig. S5), suggesting that live Pi imaging could be conducted in aerial tissues. 12

13

Live Imaging of Cytosolic Pi in Root Epidermal Cells 14

Live cell imaging using a spinning disk confocal microscope equipped with a 15

beam splitter for simultaneous dual emission fluorescence was used to test the 16

functionality of cpFLIPPi sensors for reporting changes in cytosolic Pi concentrations. 17

Untransformed plants and transgenic plants expressing eCFP and cpVenus separately 18

were imaged each day to correct for fluorescence background, spectral bleed-through 19

and cross excitation. T2 generation transgenic seedlings were grown for 5 days in 20

hydroponic medium containing 0.5 mM Pi then placed in an imaging chamber 21

containing the same medium. 22

Images for multiple epidermal cells within the root differentiation zone were 23

captured from at least six independent seedlings of each genotype to establish the steady-24

state baseline for the FRET ratio under this Pi-sufficient growth condition. The same 25

seedlings were then transferred to medium that lacked Pi and imaged again after 24 26

hours of Pi starvation. This duration of Pi starvation was chosen to avoid morphological 27

changes or cell death associated with more severe Pi deprivation (Sanchez-Calderon et 28

al., 2005). Specifically, cell viability tests with SYTOX orange (Truenit and Haseloff, 29



12

2008) consistently revealed a substantial number of non-viable cells when plants were Pi 1

starved for 72 hours (Supp. Fig. S6), but none or very few were detected when plants 2

were held in Pi-replete conditions or starved for 24 hours. Sensitivity to prolonged Pi 3

deprivation is undoubtedly a function of growth conditions and the age of the plant when 4

deprivation is initiated. 5

Separate experiments demonstrated that our Pi starvation regime was sufficient 6

to reduce the spatially-undefined Pi contents of root extracts by 19% (from 5.3 ± 0.2 7

nmol Pi/ “fed” seedling to 4.3 ± 0.2 nmol Pi/ starved seedling, mean ± SE, n=3), and was 8

also sufficient to elicit a 20% increase in secreted acid phosphatase activity (from 60 ± 4 9

mUnits/ “fed” seedling to 72 ± 5 mUnits/ starved seedling, mean ± SE, n=12), which is 10

indicative of Pi starvation (Robinson et al., 2012). Both of these changes were 11

statistically significant as determined by Student’s t-test, p<0.05). Moreover, secreted 12

acid phosphatase activities were identical in wild type and sgs3-13 plants, indicating that 13

this mutation has no obvious effect on normal responses to Pi starvation. Consequently, 14

sgs3-13 was used as the “wild-type” background for all subsequent experiments. 15

We screened transgenic lines for cpFLIPPi-80μ, -200μ, -1.6m, -4.8m, -5.3m, and 16

-6.4m under Pi-replete and 24 hr Pi starvation conditions and found that only those with 17

Kd values of 4.8 mM and greater showed a significant change in FRET ratio and in each 18

of these cases the ratio increased in response to Pi starvation, consistent with decreased 19

cytosolic Pi concentrations. Transgenic lines expressing the cpFLIPPi-6.4m sensor 20

showed the largest response and representative FRET ratio images and quantitation 21

derived from six independent seedlings are shown in Fig. 4. We attributed this response 22

to altered Pi concentrations rather than unrelated changes in the emission of one or both 23

fluorophores because no differences in FRET ratios were detected in seedlings that 24

express a high-affinity sensor, i.e., cpFLIPPi-80μ (Supp. Fig. S7A). 25

Comparisons of fluorescence intensities in cpFLIPPi-6.4m seedlings to those of 26

purified sensor protein standards imaged under identical conditions indicated that the 27

cytosolic concentrations of this sensor was only ~1 μM (Supp. Fig. S8). Consequently, 28

we suggest that the fraction of cytosolic Pi sequestered by this sensor is negligible. 29



13

To test whether FRET ratios in Pi-starved plants could be reversed by 1

replenishment with Pi we added 0.5 mM Pi to 24 hour-starved cpFLIPPi-6.4 lines and 2

monitored FRET over time (Fig. 5). No changes were detected when FRET was 3

monitored for the first few minutes. However, changes in FRET ratios indicative of 4

replenishment were detected after 1 hour, and FRET signals were nearly equivalent to 5

the initial “fed” state within 8 hours (Fig. 5). These results strongly indicate that the in 6

vivo FRET responses are specific for Pi. 7

It was possible that, within limits, the steady state concentration of Pi in the 8

cytosol is a variable continuum dependent on the magnitude of Pi supply. Alternatively, 9

if other cellular pools buffer cytosolic Pi (Mimura, 1999) then the concentration would 10

be relatively constant until buffering capacity is exceeded. To distinguish these 11

possibilities we grew transgenic cpFLIPPi-6.4 lines in hydroponic medium containing 12

0.5 mM Pi for 5 days then replaced the medium with fresh media containing the same or 13

reduced concentrations of Pi. Plants were grown in the new condition for another 24 14

hours then imaged in the same final growth medium. As shown in Fig. 6, FRET ratios 15

increased when supplied with 25 μM or less Pi, but remained nearly constant when 16

plants were supplied with Pi concentrations greater than 25 μM. FRET ratios reached a 17

maximum when plants were supplied with 10 μM Pi, but the values were not 18

significantly different from those measured with plants completely deprived of Pi. These 19

results support the idea that cytosolic Pi is buffered over a broad range of external Pi 20

concentrations, and that this buffering capacity is overwhelmed when Pi supply drops 21

below a critical threshold. 22

It was formally possible that the Pi-dependent changes in FRET ratios we 23

detected were caused by a mechanism other than FRET. Because it was not possible to 24

address this issue through analysis of the emission spectrum in vivo as was done for our 25

in vitro studies, we used partial acceptor photobleaching as an alternative approach 26

(Roszik et al., 2008). Evidence for FRET would be revealed by this method if 27

photodestruction of the acceptor cpVenus resulted in an increase in donor eCFP 28

emission due to dequenching of the donor. Moreover, the increase in eCFP emission 29



14

would be proportional to FRET efficiency, which would be dependent on Pi 1

concentration. No consistent change in eCFP emission would be expected if sensor 2

responses were unrelated to FRET. 3

We conducted partial acceptor photobleaching with each of the same plants used 4

to evaluate the effect of varied Pi supply on its accumulation in the cytosol (Fig. 6). In 5

each case, eCFP emission clearly increased as cpVenus emission diminished, indicating 6

a FRET response (Suppl. Fig. S9A). FRET efficiencies were calculated from the 7

photobleach responses as described previously (Roszik et al., 2008). The FRET 8

efficiency values calculated for each concentration of supplied Pi correlated with the 9

corresponding FRET ratios (Suppl. Fig. S9B) and thus further validated the efficacy of 10

the cpFLIPPi-6.4m sensor for monitoring cytosolic Pi. 11

12

Live imaging of Pi in Root Plastids of Wild Type and a Plastidic Pi Transport 13

Mutant 14

Nonphotosynthetic plastids import ATP from the cytosol in stoichiometric 15

exchange for stromal ADP (Reiser et al., 2004; Reinhold et al., 2007). A consequence 16

of the unbalanced phosphate moieties associated with this exchange is that Pi would 17

accumulate to deleterious levels if not balanced by export. We proposed previously 18

that the plastidic Pi transporter PHT4;2 confers this export activity in root plastids 19

(Irigoyen et al., 2011). Transport activities measured with root plastids isolated from 20

wild type and a pht4;2 mutant support this idea. However, the mutation also reduced 21

Pi import. It was possible that the import activity simply reflected the reversibility of 22

the transport process since maintenance of relevant electrochemical gradients cannot 23

be assured with isolated organelles. Alternatively, PHT4;2 may have an undefined 24

physiological role in Pi import. 25

To directly assess the role of PHT4;2 in plastidic Pi transport within the context 26

of an intact plant we introduced the plastid-targeted version of the cpFLIPPi-6.4m sensor 27

into the pht4;2 background then compared FRET ratios in this line to those of the same 28

sensor in wild type. Plants were grown for 5 days in medium containing 0.5 mM Pi to 29



15

establish uniform growth then imaged in the same medium. FRET ratios were 1

significantly lower in mutant plastids, which is consistent with increased accumulation 2

of Pi. Representative ratio images and quantitation from six independent seedlings of 3

each genotype are shown in Fig. 7. The FRET ratio images were merged with DIC 4

images to distinguish individual cells, and this revealed that there is little variation in the 5

amounts of Pi accumulated in plastids within the same cell or in different epidermal cells 6

(Fig. 7A). The differences in FRET ratios can be attributed to altered Pi concentrations 7

rather than unrelated effects of the pht4;2 mutation because no differences in FRET 8

ratios were detected when the high-affinity plastid cpFLIPPi-80μ sensor was used in 9

these same genotypes (Supp. Fig. S7B). These results strongly support the hypothesis 10

that the physiologically-relevant role of PHT4;2 is Pi export from root plastids. 11

12

DISCUSSION 13

Despite variable and often scarce supplies of Pi from soils, plants must distribute 14

appropriate amounts of Pi to each cell and subcellular compartment to sustain essential 15

metabolic activities. Because its concentrations in different locations can vary widely 16

and in some cases may also serve as a signal of Pi sufficiency (Carswell et al., 1996; 17

Ticconi et al., 2001; Varadarajan et al., 2002; Kanno et al., 2012), dynamic assessment 18

of these concentrations in live plants is critical for understanding the complex 19

mechanisms underlying Pi homeostasis. Nonaqueous fractionation and 31P-NMR have 20

been used to probe Pi contents of some cell compartments, but these methods lack 21

cellular resolution and have no or limited temporal resolution (Rebeille et al., 1984; 22

Sharkey and Vanderveer, 1989; Stitt et al., 1989; Pratt et al., 2009; Gout et al., 2011). 23

Therefore, to achieve the necessary spatial and temporal resolution we chose to adapt for 24

use in plants a genetically encoded Pi sensor (FLIPPi) that had been used previously to 25

track Pi in cultured animal cells (Gu et al., 2006). 26

Our efforts to adapt FLIPPi sensors for use in plants included modification of the 27

sensor proteins first to enhance dynamic range and then to optimize affinity. To enhance 28

dynamic range, i.e., the maximum change in Pi-dependent FRET, we used a strategy that 29



16

had proven successful for improvement of yellow cameleon Ca2+ sensors (Nagai et al., 1

2004). Specifically, we substituted the eYFP component of a FLIPPi sensor with a 2

circularly permuted form of Venus (cpVenus). The resulting circularly permuted FLIPPi 3

(cpFLIPPi) sensor exhibited increased FRET even in the absence of Pi. More 4

importantly, its dynamic range was more than twice that of the parent sensor. This 5

enhancement is presumably a function of the relative orientation of the two 6

chromophores in the sensor rather than their distance because the chromophores remain 7

equally separated by the Pi binding protein (Nagai et al., 2004). Regardless, it is unlikely 8

that cpVenus and eYFP interact differently with the Pi binding protein component of the 9

sensor because binding affinity was unaffected. 10

Most estimates of cellular Pi concentrations in Pi-replete plants range from 2.5 to 11

10 mM and are derived from enzyme kinetic parameters, nonaqueous fractionation and 12 31P-NMR (Rebeille et al., 1984; Stitt et al., 1988; Sharkey and Vanderveer, 1989; Lee 13

and Ratcliffe, 1993; Copeland and Zammit, 1994; Gout et al., 2011; Tiessen et al., 14

2012), although cytosolic concentrations as low as 60 to 80 μM have also been reported 15

(Pratt et al., 2009). We used these estimates as a guide for selecting cpFLIPPi affinity 16

mutants with a goal of encompassing the entire range. Our mutagenesis strategy 17

involved a novel use of statistical coupling analysis (SCA) (Lockless and Ranganathan, 18

1999; Socolich et al., 2005) as a means to enhance the efficiency of the screen. The 19

premise of SCA is that coupling represents an evolutionary constraint for structural or 20

functional properties, e.g., ligand binding, protein-protein interactions and allosteric 21

sites. We conducted SCA with a set of 1,020 conserved Pi binding proteins and 22

identified a six amino acid region that contained coupled residues that also map near the 23

predicted Pi binding site. By limiting our mutagenesis to alter only this six amino acid 24

region rather than exploring the entire 340 amino acids of the Pi binding protein 25

included in the cpFLIPPi sensor, we reduced the scope of mutagenesis numerically and 26

also greatly biased our screen for those likely to alter Pi binding properties. 27

The mutant screen was highly efficient with nearly 80% of the individuals 28

screened showing altered affinity. Eight of these mutants were selected for further 29



17

analysis. Each of these were highly specific for Pi and had Kd values for Pi binding that 1

ranged from 0.08 to 11 mM, which spanned the entire target range. Although the in vitro 2

binding assays were conducted in a medium that approximated cellular viscosity and 3

primary ionic composition (Messerli and Robinson, 1998; Feijo et al., 1999), we 4

recognize that the binding affinities may differ in vivo. Nevertheless, the large (>100-5

fold) concentration range collectively probed by the cpFLIPPi sensors was more than 6

sufficient to monitor relative changes in the Pi contents of cytosol and plastid stroma of 7

Arabidopsis root epidermal cells and are likely to also be useful for monitoring Pi 8

contents in other cells and other cell compartments with near-neutral pH. 9

The cpFLIPPi sensors yielded weak or unstable fluorescent signals when 10

expressed in wild-type plants suggesting that they were subject to transgene silencing. 11

Silencing was also reported for related FRET-based sensors for glucose and glutamine, 12

and in these cases silencing was eliminated when the sensors were expressed in siRNA-13

deficient mutants sgs3 and rdr6 (Deuschle et al., 2006; Chaudhuri et al., 2008; Yang et 14

al., 2010). It is possible that some aspect of the ligand-binding portions of these sensors 15

promotes their silencing because a related Ca2+ sensor that contains the same 16

fluorophores as cpFLIPPi and is expressed from the same Arabidopsis UBQ10 promoter 17

is not subject to silencing (Krebs et al., 2012). Regardless, we attained strong, stable 18

fluorescence when cpFLIPPi sensors were expressed in the sgs3-13 background. It 19

should be noted that although rdr6 mutations are more effective at eliminating transgene 20

silencing than sgs3 mutations (Deuschle et al., 2006), we chose to use sgs3-13 for our 21

studies because, unlike rdr6 mutations, it had little effect on plant growth and 22

morphology. Moreover, secreted acid phosphatase activities measured in Pi-sufficient 23

and Pi-starved sgs3-13 seedlings were equivalent to those of wild-type seedlings grown 24

under the same conditions suggesting that this mutation has no significant effect on 25

normal responses to Pi starvation. 26

Analysis of root extracts revealed that depriving 5-day old seedlings of Pi for 24 27

hours was sufficient to reduce free, cellular- and subcellular-undefined Pi contents by 28

~19%. Live imaging of cpFLIPPi sensors allowed us to unambiguously monitor the 29



18

fraction attributed to cytosolic Pi contents of epidermal cells. A significant change in 1

FRET ratio was detected only for sensors with Kd values of 4.8 mM or greater, which 2

suggests that sensors with higher affinity remained effectively saturated under this 3

relatively mild starvation regime. Transgenic lines with the cpFLIPPi-6.4m sensor 4

yielded the largest FRET response upon Pi deprivation, which may reflect an optimal 5

pairing of the binding affinity of this sensor with cytosolic Pi concentrations. 6

Multiple lines of evidence indicate that the changes in FRET ratios we measured 7

in epidermal cells represent genuine Pi-dependent responses. First, FRET responses 8

elicited by Pi deprivation were completely reversed by Pi replenishment. Second, high-9

affinity sensors exhibited no detectable FRET responses suggesting little or no changes 10

in Pi-independent factors that could influence sensor properties. And third, Pi-dependent 11

FRET efficiencies determined from acceptor photobleaching experiments directly 12

correlated with the corresponding FRET ratios. Consequently, we conclude that the 13

cpFLIPPi-6.4m sensor reports changes in cytosolic Pi concentrations with high 14

specificity. Our results also suggest that the concentration of Pi in the cytosol is 15

relatively constant despite large differences in the amount of Pi supplied in the growth 16

medium, presumably due to buffering by vacuolar pools (Mimura, 1999). This buffering, 17

however, can be overwhelmed when Pi supply falls below a critical threshold. For our 18

growth and assay conditions, this threshold was ~25 μM. 19

In addition to cytosolic Pi, the cpFLIPPi-6.4 sensor also enabled monitoring of Pi 20

at the subcellular level. We proposed previously that the physiologically relevant role of 21

PHT4;2, a plastidic Pi transporter, was export of Pi from root plastids (Irigoyen et al., 22

2011). This hypothesis was based on transport assays conducted with isolated root 23

plastids, but the assays also revealed an import activity, which we viewed as a 24

consequence of transport reversibility under our assay conditions. However, it was 25

formally possible that Pi import was the relevant biological activity. Ratiometric 26

imaging of a plastid-targeted version of cpFLIPPi-6.4m expressed in wild-type and 27

pht4;2 plants distinguished these possibilities in the context of a live plant and thereby 28

confirmed that Pi export is the physiologically relevant role for PHT4;2. 29



19

1

CONCLUSIONS 2

The current series of cpFLIPPi sensors provide the necessary spatial and 3

temporal resolution to follow critical aspects of Pi metabolism in live plants. One of 4

these, cpFLIPPi-6.4m, was particularly useful because it detected reduced cytosolic Pi 5

concentrations in response to varied external supply and also detected a mutation-6

induced increase in Pi accumulation within an organelle. These sensor proteins are also 7

likely to be suitable for studies in microbial and animal cell systems, and the approaches 8

described here for their optimization may be broadly applicable to other genetically 9

encoded sensors. Future efforts to target cpFLIPPi sensors to additional subcellular 10

compartments and further engineering to enhance signal responses in acidic 11

environments will provide valuable insight into the mechanisms underlying Pi 12

metabolism as well as the sensing and signaling of Pi status. 13

14

15

MATERIALS AND METHODS 16

Plasmids 17

We obtained FLIPPi plasmid clones pRSET FLIPPi-200μ and pRSET FLIPPi-18

30m (Gu et al., 2006) from Addgene (www.Addgene.com). To replace the eYFP coding 19

region of pRSET FLIPPi-200μ with a circularly permuted version of Venus, we first 20

amplified the cp173 coding region from the YC3.60 Ca2+ sensor (Nagai et al., 2004). We 21

then used PCR and the In-Fusion (Clontech) cloning kit to introduce this sequence into 22

the same region occupied by the eYFP coding sequence in the pRSET FLIPPi-200μ 23

plasmid to generate pRSET cpFLIPPi-200μ. 24

For cpFLIPPi sensor expression in plants, a 2455 bp Hind III-BamHI segment of 25

the desired pRSET cpFLIPPi clone was ligated into pCN-UBQ10, a modified 26

pCAMBIA3300 plasmid. The modifications included insertion of the Arabidopsis 27

UBQ10 promoter (Norris et al., 1993) to direct expression of the sensor gene and 28

replacement of the 35S promoter located upstream of the phosphinothricin resistance 29



20

gene with the Nos promoter of pBI121. A plastid-targeted variant of each clone was 1

constructed by inserting a 237 bp fragment encoding the 79 amino acid RbcS chloroplast 2

transit peptide (Lee et al., 2006) in frame at the 5’ end of the sensor gene. 3

4

Construction of a cpFLIPPi affinity mutant library 5

We used statistical coupling analysis (SCA) (Lockless and Ranganathan, 1999; 6

Socolich et al., 2005) as a guide to generate a sensor mutant library biased for altered 7

affinity. Sequences were collected from the nr database (as of 5/5/2010) using PSI-8

BLAST (Altschul et al., 1997). The Escherichia coli PiBP protein (GI:157829674) was 9

used as the query for five iterations of PSI-BLAST. Sequences with an e-score<1e-10 10

were aligned using MUSCLE (Edgar, 2004). Identical sequences were eliminated and 11

the alignment of the remaining 1020 sequences was used for SCA. SCA was performed 12

as previously described (Lockless and Ranganathan, 1999; Socolich et al., 2005) to 13

identify the set of positions that co-evolve with one another. Two rounds of hierarchical 14

clustering were performed to yield a self-consistent set of positions that statistically co-15

vary with each other. PiBPs have one self-consistent cluster consisting of 43 amino acids 16

(13% of protein), which was mapped onto the structure of the E. coli PiBP using PyMol 17

(Suppl. Fig. S3). 18

A stretch of six amino acids (positions 18-23) within the 43 amino acid cluster 19

was identified as a high-value mutational target based on the following observations: 1) 20

four of the positions (#18, 19, 21 and 23) co-evolve with one another, 2) one position 21

(#20) is highly conserved, 3) they are located near the Pi binding site, and 4) the 22

positions could be simultaneously mutated with one set of oligonucleotides. The 23

mutagenesis oligonucleotides were designed such that each position was mutated to the 24

three most frequently observed amino acids in the alignment that are found in at least 9% 25

of sequences. In practice, this means that more than the desired amino acids are possible 26

at some positions since the nucleotides required to encoded the target amino acids can 27

also encode other amino acids. For example, alanine (GCT codon), aspartate (GAT 28

codon) and serine (TCT codon) are needed at position 19, but the nucleotides required in 29



21

the oligonucleotide at each position can also encode for tyrosine (TAT codon). 1

The pRSET cpFLIPPi-200μ plasmid was used as template for PCR mutagenesis. 2

The mutagenesis oligonucleotides 5’- 3

ccaaaccgtgcaaatctccRBCKMTgggcKCGDMCDYCgcggctcctttgctgcaacgttg and 5’- 4

gttgcagcaaaggagccgcGRHGKHCGMgccAKMGVYggagatttgcacggtttgggctt introduce 5

ca. 1728 possible amino acid combinations for the six amino acid (positions 18-23) 6

target region. The amplicon population was restriction digested then ligated into pRSET 7

cpFLIPPi-200μreplacing the original sequence. A portion of this mixed ligation product 8

was introduced into E. coli DH5α cells and >10,000 colonies were pooled for bulk 9

plasmid isolation. The isolated plasmids collectively represent the affinity mutant 10

library. 11

12

cpFLIPPi affinity mutant screen 13

Portions (0.1 ng) of the sensor affinity mutant plasmid library were introduced 14

into E. coli BL21 (DE3) cells. The pRSET cpFLIPPi-200μ plasmid served as a positive 15

control and pUC19 served as a negative control for each transformation and subsequent 16

screen. Individual colonies were transferred to a 96-well plate containing 45 μL LB 17

medium in each well and mixed thoroughly. Aliquots of 20 µL were transferred to 18

duplicate 96-well deep well plates containing 1.5 mL LB medium then covered with a 19

gas-permeable seal. Plates were shaken at 300 rpm for 16 hours at 25°C. Glycerol stocks 20

were prepared from 0.1 mL aliquots of each culture in a separate 96-well plate and 21

stored at -80°C. The growth plates were centrifuged for 30 min to pellet cells then the 22

supernatants were removed by vacuum aspiration. Cell pellets were washed twice with 23

20 mM Tris-HCl, pH 7.5 then frozen at -20°C. To lyse cells, 0.35 mL lysis solution (1x 24

BugBuster (Novagen), 20 mM Tris-HCl pH 7.5, 150 mM K-gluconate, 3 μL 25

Lysonase/mL) was added to each well, mixed thoroughly then incubated at room temp 26

for 20 min with mixing every 5 minutes. Plates were then centrifuged for 10 min at 2800 27

rpm, 20°C to remove cell debris. 28



22

Fluorescence assays were conducted with lysates in 96-well black plates. We 1

transferred 20 μL aliquots of each lysate to eight wells containing 200 μL pseudocytosol 2

medium (100 mM K-gluconate, 30 mM NaCl, 25 mM MES, 25 mM HEPES, 40% 3

sucrose, 1 mg/mL BSA, pH 7.5) (Messerli and Robinson, 1998; Feijo et al., 1999) and 4

varied amounts of Pi (0 to 30 mM). Fluorescence was measured using a microplate 5

reader (Synergy HT) with excitation 420/27 nm and emission 485/20 and 540/25 nm. 6

Fluorescence from direct excitation of cpVenus (500/20 nm) was also monitored with 7

emission at 540/25 nm. The Kd value for each sensor was estimated by fitting data to a 8

single-site binding isotherm as described (Gu et al., 2006). 9

10

In vitro characterization of cpFLIPPi proteins 11

Selected cpFLIPPi mutant clones were re-introduced into E. coli BL21 (DE3) 12

cells, and single colonies were cultured in 100 mL medium then harvested and lysed 13

essentially as described above for the mutant screen. The expressed sensor proteins were 14

purified from cell lysates using His-affinity chromatography with elution in 20 mM Tris-15

HCl pH 7.5, 150 mM K-gluconate, 400 mM imidazole. The eluent was dialyzed 16

overnight at 4°C against 20 mM Tris-HCl pH 7.5 then transferred to a new tube and 17

stored at 4°C. The purified sensors could be stored for at least one month without 18

measurable change in fluorescence or binding properties. Kd values were determined as 19

described above for the mutant screen, but twelve Pi concentrations were used to 20

generate binding curves. Assays for binding affinity, specificity and pH sensitivity were 21

all conducted with at least three independent protein preparations. 22

23

Generation of transgenic Arabidopsis plants 24

Sensor transgenes were introduced into Arabidopsis plants by floral dip 25

transformation (Clough and Bent, 1998). 26

27

Subcellular localization of sensor proteins 28



23

Protoplasts were prepared as described (Yoo et al., 2007) from leaves of 5-week-1

old transgenic plants expressing cpFLIPPi sensors engineered with and without an N-2

terminal RbcS chloroplast transit peptide. Protoplasts were imaged with a BioRad ES 3

confocal laser scanning microscope equipped with a 63x (numerical aperture, 1.2) water-4

immersion objective (515 nm excitation, 540 nm emission for direct excitation of the 5

cpVenus sensor component, and 488 nm excitation, 680 nm emission for chlorophyll 6

autofluorescence. 7

8

Live imaging of sensors in plants 9

Transgenic plants were grown in individual wells of 96-well plates with 0.3 mL 10

of half-strength MS medium containing 1% sucrose and unless indicated otherwise, 0.5 11

mM Pi. The plates were incubated in a growth chamber (60% relative humidity, 21°C, 12

90 μmol m-2 s-1 light intensity, 16 hour photoperiod). After 5 days the seedlings were 13

either imaged or transferred to fresh medium as indicated. For imaging, seedlings were 14

placed in a custom-made chamber (http://microscopy.tamu.edu/lab-protocols/light-15

microscopy-protocols.html) containing the same growth medium. Seedlings were 16

covered with a small square of nylon mesh then gently immobilized with a glass weight. 17

A similar procedure was described recently (Kruger et al, 2013). Seedlings were 18

analyzed for FRET using an inverted Olympus IX81 laser spinning disk confocal 19

microscope equipped with an Andor Opti-splitter to allow simultaneous dual emission. 20

Root epidermal cells were viewed with a 40x (1.3 n.a.) oil-immersion objective (445 nm 21

excitation for eCFP and 483/32 and 542/27 nm emission wavelengths for eCFP and 22

FRET-derived cpVenus, respectively. Laser intensity and exposure settings were kept 23

constant. For acceptor photobleaching experiments, a 515 nm excitation laser was used 24

with maximum laser intensity. Untransformed seedlings and transgenic seedlings 25

expressing eCFP alone and cpVenus separately were imaged each day to allow 26

correction for background fluorescence, spectral bleedthrough and cross excitation. 27

All image analyses were done using ImageJ software. Mean intensity values for 28

untransformed seedlings were used to establish background fluorescence values, and 29



24

these were subtracted from all sensor images before conducting further analyses. For 1

analysis of cytosol-localized sensors, mean intensity values for eCFP and cpVenus were 2

measured for selected regions of interests (ROIs), which typically included 3-4 cells for 3

each field of view. For analysis of plastid-localized sensors, a uniform threshold was 4

applied to the eCFP and FRET-cpVenus images, and mean intensity values above the 5

threshold were used to calculate sensitized FRET ratios. FRET efficiencies (E values) 6

were calculated from partial acceptor photobleaching images as described (Roszik et al., 7

2008). Briefly, mean intensity values for eCFP (donor) and directly excited cpVenus 8

(acceptor) for each sample were recorded prior to photobleach and again for at least four 9

photobleach time points. These values were corrected for spectral bleedthrough and 10

cross-excitation then used to calculate mean FRET efficiencies. An ImageJ plugin, 11

AccPbFRET, is available to automate much of this analysis (Roszik et al., 2008). 12

13

Colorimetric Pi assay 14

Plants were grown exactly as for live imaging with and without a final 24 hour 15

Pi deprivation. Roots were harvested and pooled in groups of 15. Pi contents were 16

determined using a phosphomolybdate colorimetric assay as described previously 17

(Ames, 1966; Versaw and Harrison, 2002). 18

19

Secreted acid phosphatase assay 20

Plants were grown exactly as for live imaging with and without a final 24 hour 21

Pi deprivation. Individual seedlings were then transferred to 0.3 mL fresh medium 22

containing no Pi. Secreted acid phosphatase activity in the medium was measured after 23

30 minutes using p-nitrophenol phosphate as described (Robinson et al., 2012) except 24

seedlings were removed from the assay mixture immediately prior to termination with 25

1M sodium carbonate. One unit of phosphatase activity is defined as 1 μmol p-26

nitrophenol liberated per minute. 27

28

Upon request, all novel materials described in this publication will be made 29



25

available in a timely manner for non-commercial research purposes, subject to the 1

requisite permission from any third-party owners of all or parts of the material. 2

Obtaining any permissions will be the responsibility of the requestor. 3

4

SUPPLEMENTAL MATERIAL 5

Supplemental Figure S1. Emission spectrum of cpFLIPPi-200μ in pseudcytosol 6

medium containing 0 or 10 mM Pi. 7

8

Supplemental Figure S2. Analysis to identify residues that co-evolve in the PiBP 9

protein family. 10

11

Supplemental Figure S3. Mapping of co-evolving and mutated residues onto the PiBP 12

structure (pdb1A40). 13

14

Supplemental Figure S4. Specificity of cpFLIPPi-6.4m. 15

16

Supplemental Figure S5. Differential targeting oof cpFLIPPi-6.4m in palisade 17

mesophyll cells of intact Arabidopsis leaves. 18

19

Supplemental Figure S6. SYTOX orange staining shows non-viable cells in roots after 20

72 hr Pi starvation. 21

22

Supplemental Figure S7. Live imaging of cytosolic and plastidic Pi in root epidermal 23

cells using a high-affinity Pi sensor, cpFLIPPi-80μ. 24

25

Supplemental Figure S8. Estimates of cpFLIPPi-6.4m protein concentrations in the 26

cytosol of root epidermal cells. 27

28

Supplemental Figure S9. Pi-dependent FRET efficiencies determined from partial 29



26

acceptor photobleaching of cytosolic cpFLIPPi-6.4m correlate with FRET ratio values. 1

2

ACKNOWLEDGEMENTS 3

We thank Shian Liu for assistance with analysis of the mutant library and Tom 4

McKnight for critical reading of this manuscript. 5

6

7

8



27

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Vance CP, Uhde-Stone C, Allan DL (2003) Phosphorus acquisition and use: critical 3 adaptations by plants for securing a nonrenewable resource. New Phytol 157: 4 423-447 5

Varadarajan DK, Karthikeyan AS, Matilda PD, Raghothama KG (2002) Phosphite, 6 an analog of phosphate, suppresses the coordinated expression of genes under 7 phosphate starvation. Plant Physiol 129: 1232-1240 8

Versaw WK, Harrison MJ (2002) A chloroplast phosphate transporter, PHT2;1, 9 influences allocation of phosphate within the plant and phosphate-starvation 10 responses. Plant Cell 14: 1751-1766 11

Walker DA, Sivak MN (1986) Photosynthesis and phosphate: a cellular affair? Trends 12 Biochem Sci 11: 176-179 13

Yang H, Bogner M, Stierhof YD, Ludewig U (2010) H+-independent glutamine 14 transport in plant root tips. PLoS One 5: e8917 15

Yoo SD, Cho YH, Sheen J (2007) Arabidopsis mesophyll protoplasts: a versatile cell 16 system for transient gene expression analysis. Nat Protoc 2: 1565-1572 17

Zhang Z, Liao H, Lucas WJ (2014) Molecular mechanisms underlying phosphate 18 sensing, signaling, and adaptation in plants. J Integr Plant Biol 56: 192-220 19

20

21

22



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Figure legends 1

2

Figure 1. Pi-dependent changes in FRET ratios of purified Pi sensors. A, Schematic of 3

Pi sensor indicating high FRET in absence of Pi and reduced FRET when bound to Pi. 4

Acceptor is eYFP for FLIPPi sensors and cpVenus for cpFLIPPi sensors. PiBP, Pi 5

binding protein. B, Values shown are means ± SE for three independent protein 6

preparations. Error bars smaller than the symbols are not shown. 7

8

Figure 2. Effect of pH on Pi-dependent FRET responses. Pi binding of cpFLIPPi-6.4m 9

was tested in pseudocytosol mdium adjusted to pH 7.0, 7.5 and 8.0, and Pi was adjusted 10

to the same pH. Binding affinity (Kd) values were estimated from nonlinear regression 11

analysis. Plotted values are means ± SE for three independent protein preparations. Error 12

bars smaller than the symbols are not shown. 13

14

Figure 3. Differential targeting of cpFLIPPi-6.4m. Protoplasts were isolated from leaves 15

of stably transformed Arabidopsis plants expressing cpFLIPPi-6.4m with or without an 16

N-terminal chloroplast transit peptide. Confocal microscoy was conducted using settings 17

for cpVenus excitation and emission (green) and chlorophyll autofluorescence 18

(magenta). Scale bar, 10 μm. 19

20

Figure 4. Live imaging of cytosolic Pi in root epidermal cells. A, Representative FRET 21

ratio images of cpFLIPPi-6.4m stably expressed in seedlings with sufficient Pi (control, 22

left), and after 24 hr Pi starvation (right). The same color scale was applied to ratio 23

values for both images. Size bar, 10 μm. B, Plotted FRET ratio values are means ± SE 24

for six independent seedlings. The difference between the two conditions is significant, 25

p<0.05, Student’s t-test. 26

27

Figure 5. Replenishment of cytosolic Pi after 24 hr starvation. Seedlings expressing 28

cpFLIPPi-6.4m were grown with sufficient Pi (0.5 mM, +Pi), starved for 24 hr (-Pi) then 29



33

Pi was added to 0.5 mM and FRET ratios were monitored over time. The plotted values 1

are means ± SE for six independent seedlings. The first time point shown after 2

replenishment is 1 hr. * indicates significant difference (p<0.05, Student’s t-test) from 3

the –Pi treatment. 4

5

Figure 6. Effect of Pi supply on cytosolic contents of root epidermal cells. Seedlings 6

expressing cpFLIPPi-6.4m were grown with a sufficient supply of Pi (500 μM) then 7

transferred to medium containing the same or less Pi as indicated and grown for an 8

additional 24 hr before imaging. Plotted FRET ratio values are means ± SE for at least 9

six independent seedlings. * indicates significant difference (p<0.05, Student’s t-test) 10

from the 500 μM treatment. 11

12

Figure 7. Live imaging of Pi in plastids of root epidermal cells. A, Representative FRET 13

ratio images for cpFLIPPi-6.4m stably expressed in wild-type and pht4;2 seedlings. The 14

images were merged with the corresponding DIC images to distinguish cell boundaries. 15

The same color scale was applied to ratio values for both images. Size bar, 5 μm. B, 16

Plotted FRET ratio values are means ± SE for six independent seedlings. The difference 17

between the two genotypes is significant, p<0.05, Student’s t-test. 18

19

20



34

Table I. Pi sensor affinity mutants 1 The cpFLIPPi-200μ sensor plasmid was subjected to a PCR-based mutagenesis. The sequences 2 shown correspond to the mutated region of the sensor. Pi binding constants were determined in vitro using 3 three independent protein preparations for each sensor. ∆Rmax indicates the maximum change in FRET 4 ratio between Pi-free and Pi-saturated sensor. 5 6

Sensor Kd (mM) ∆Rmax DNA sequence of PiBP cds Protein sequence cpFLIPPi-80μ 0.08 -1.06 49 TCCGtcGCtGGCtCGACCaccGCG 17 SVAGSTTA

cpFLIPPi-200μ 0.2 -1.08 49 TCCGGGGCGGGCGCGACCTTTGCG 17 SGAGATFA cpFLIPPi-1.6m 1.6 -0.91 49 TCCGccGatGGCtCGACCaTcGCG 17 SADGSTIA cpFLIPPi-3.5m 3.5 -0.43 49 TCCGGcGCtGGCtCGAaGTcTGCG 17 SGAGSKSA cpFLIPPi-4.8m 4.8 -0.76 49 TCCaccGatGGCGCGACCgccGCG 17 STDGATAA cpFLIPPi-5.3m 5.3 -0.71 49 TCCaGcGatGGCtCGtCCgtcGCG 17 SSDGSSVA cpFLIPPi-6.4m 6.4 -0.75 49 TCCGGGGCGGGCGCGgCCTTTGCG 17 SGAGAAFA cpFLIPPi-11m 11.0 -0.45 49 TCCatctCtGGCtCGtCCaTcGCG 17 SISGSSIA

7 8



Figure 1. Pi-dependent changes in FRET ratios of purified Pi sensors. A, Schematic of Pi sensor indicating high FRET in absence of Pi and reduced FRET when bound to Pi. Acceptor is eYFP for FLIPPi sensors and cpVenus for cpFLIPPi sensors. PiBP, Pi binding protein. B, Values shown are means ± SE for three independent protein preparations. Error bars smaller than the symbols are not shown.

FRET

acceptor eCFP

PiBP

Pi acceptor eCFP

PiBP A

0.5

1

1.5

2

2.5

0 4 8 12 16 20

FLIPPi-200µ

cpFLIPPi-200µ

FRET

ratio

(cpV

enus

/eC

FP)

Pi (mM)

B



Figure 2. Effect of pH on Pi-dependent FRET responses. Pi binding of cpFLIPPi-6.4m was tested in pseudocytosol medium adjusted to pH 7.0, 7.5 and 8.0, and Pi was adjusted to the same pH. Binding affinity (Kd) values were estimated from nonlinear regression analysis. Plotted values are means ± SE for three independent protein preparations. Error bars smaller than the symbols are not shown.

1.2

1.4

1.6

1.8

2.0

0 5 10 15 20

FRET

ratio

(cpV

enus

/eC

FP)

Pi (mM)

Kd (mM)

pH 7.0 4.9 ± 0.3

pH 7.5 6.4 ± 0.4

pH 8.0 9.6 ± 0.5



Figure 3. Differential targeting of cpFLIPPi-6.4m. Protoplasts were isolated from leaves of stably transformed Arabidopsis plants expressing cpFLIPPi-6.4m with or without an N-terminal chloroplast transit peptide. Confocal microscopy was conducted using settings for cpVenus excitation and emission (green) and chlorophyll autofluorescence (magenta). Scale bar, 10 µm.

Plas

tid-lo

caliz

ed

cpFL

IPPi

-6.4

m

cpVenus Chlorophyll Merged

Cyt

osol

-loca

lized

cp

FLIP

Pi-6

.4m



Pi starved 24 hr Control 1.0 1.2 1.4 1.6 1.8 2.0 2.2

FRET

ratio

(cpV

enus

/eC

FP)

Figure 4. Live imaging of cytosolic Pi in root epidermal cells. A, Representative FRET ratio images of cpFLIPPi-6.4m stably expressed in seedlings with sufficient Pi (control, left), and after 24 hr Pi starvation (right). The same color scale was applied to ratio values for both images. Size bar, 10 µm. B, Plotted FRET ratio values are means ± SE for six independent seedlings. The difference between the two conditions is significant, p<0.05, Student’s t-test.

B

Pi starved 24 hr Control A



Figure 5. Replenishment of cytosolic Pi after 24 hr starvation. Seedlings expressing cpFLIPPi-6.4m were grown with sufficient Pi (0.5 mM, +Pi), starved for Pi for 24 hr (-Pi) then Pi was added to 0.5 mM and FRET ratios were monitored over time. The plotted values are means ± SE for six independent seedlings. The first time point shown after replenishment is 1 hr. * indicates significant difference (p<0.05, Student’s t-test) from the -Pi treatment.

1

1.2

1.4

1.6

1.8

2.0

2.2

+Pi -Pi 4 8 12 16 20 24

FRET

ratio

(cpV

enus

/eC

FP)

Hours after Pi replenishment

* *

* *



Figure 6. Effect of Pi supply on cytosolic contents of root epidermal cells. Seedlings expressing cpFLIPPi-6.4m were grown with a sufficient supply of Pi (500 µM) then transferred to medium containing the same or less Pi as indicated and grown for an additional 24 hr before imaging. Plotted FRET ratio values are means ± SE for six independent seedlings. * indicates significant difference (p<0.05, Student’s t-test) from the 500 µM treatment.

1.0

1.2

1.4

1.6

1.8

2.0

2.2

0 10 25 50 100 500

FRET

ratio

(cpV

enus

/eC

FP)

Pi in growth medium (µM)

* *

*



WT pht4;2 A

0.0

0.5

1.0

1.5

2.0

2.5

WT pht4;2

FRET

ratio

(cpV

enus

/eC

FP)

B

Figure 7. Live imaging of Pi in plastids of root epidermal cells. A, Representative FRET ratio images for cpFLIPPi-6.4m stably expressed in wild-type and pht4;2 seedlings. The images were merged with the corresponding DIC images to distinguish cell boundaries. The same color scale was applied to ratio values for both images. Size bar, 5 µm. B, Plotted FRET ratio values are means ± SE for six independent seedlings. The difference between the two genotypes is significant, p<0.05, Student’s t-test.



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