**FIGURE 1 | Energy migration pathway in chloroplasts can be visualized by spatially resolved fluorescence excitation spectroscopy (FExS). (A)** Energy migration scheme in chloroplasts. S1 to S3 indicate states in the LHC energy transfer cascade, from which excitation energy is transferred downstream to the reaction center (S4). **(B)** Representative confocal fluorescence excitation spectrum of an *A. thaliana* chloroplast recorded by sweeping the excitation wavelength while recording the fluorescence intensity at 680 nm. **(C)** Typical chloroplast fluorescence emission spectrum of an *A. thaliana* chloroplast excited at 633 nm.

## **METHODS**

## **PLANT MATERIAL AND GROWTH CONDITIONS**

Arabidopsis (*Arabidopsis thaliana*) ecotype Columbia-0 (wild type, wt) and transgenic plants overexpressing the maize photosynthetic NADP-malic enzyme (MEm2 and MEm5 Fahnenstich et al., 2007) were grown in pots containing three parts of soil (Gebr. Patzer KG, Sinntal-Jossa, Germany) and one part of vermiculite (Basalt Feuerfest, Linz, Germany) at 22◦C day temperature under long days (LD, 16 h light/8 h darkness) or short days (SD, 8 h light/16 h darkness). The light intensity (photosynthetic active radiation) was 110μmol quanta m−2s −1. Maize plants were grown in soil with temperatures ranging from 23 to 26◦C and a 16 h light/8 h darkness cycle at a photon flux density of 350μmol quanta m−2s −1. Mature (12 week-old plants) and immature (2 week-old seedlings) maize plants were used for the measurements.

## **STATISTICAL ANALYSIS OF ROOM TEMPERATURE EMISSION SPECTRA (SART)**

Data acquisition for SART was accomplished by initially recording a fluorescence intensity map of the plant tissue to identify the chloroplasts using a custom-built confocal fluorescence microscope based on a Zeiss Axiovert 135TV (Carl Zeiss, Germany, Detector: PDM avalanche photodiode, Picoquant, Germany). The custom-built setup was equipped with a 633 nm excitation source (PL-610, Polytec, Germany) and a raster scanning stage (P517, Physik instrumente, Germany). Fluorescence emission spectra with diffraction-limited spatial resolution were acquired Peter et al. *In vivo* chloroplast fluorescence spectroscopy

from chloroplasts to determine the relative contributions of PSI and PSII to the spectral signature using a polychromator spectrograph (Acton 300i, Roper Scientific, US, Pixis 100B, Princeton Instruments, US). To reduce the data volume, this spectral shape was analyzed by studying the intensity ratio at characteristic wavelengths. The PSI to PSII intensity ratio was directly deduced by calculating the intensity ratio λemPSI(730 nm):λemPSII(680 nm). To account for individual differences in the composition within and among single chloroplasts, we recorded five spectra per chloroplast and averaged the PSI/PSII intensity ratio of numerous chloroplasts (*n* = 140–200). Based on this statistical analysis of chloroplast emission spectra at room temperature (SART), the average PSI/PSII intensity ratio of living plants was determined as a function of external conditions.

## **FLUORESCENCE EXCITATION SPECTRO-MICROSCOPY (FExS)**

To record fluorescence excitation spectra with a diffraction limited spatial resolution, our custom-built confocal microscope (details above) was combined with a supercontinuum laser (SC-400pp, Fianium, UK) as an excitation source. To sequentially scan the excitation wavelength, the supercontinuum source was coupled into a grating monochromator (Acton 300i, Roper Scientific, US), which was synchronized with the data acquisition by a custom-built software protocol. The fluorescence emission was recorded in a spectral window between 655 and 685 nm, matching the fluorescence emission of the PS II reaction center. Excitation spectra were recorded by sweeping the excitation wavelength from 500 to 620 nm, covering the absorption of the LHC pigments while recording the fluorescence emission. Using this fluorescence excitation spectroscopy (FExS) arrangement, intense fluorescence emission is observed for excitation waengths in resonance with the absorbance of one of the LHC pigments. Simultaneously, the energy migration pathway is intact so that the excitation energy is funneled downstream to the reaction center. Accordingly, the intensity of the distinct bands is proportional to the relative amount of energy guided to the reaction center (**Figure 1**). A detailed description of the experimental realization by the combination of confocal microscopy with a feedbackcontrolled supercontinuum laser source is given elsewhere (Blum et al., 2011).

## **RESULTS**

## **USING STATISTICAL ANALYSIS OF ROOM TEMPERATURE EMISSION SPECTRA (SART) TO MONITOR THE PSI/PSII INTENSITY RATIO OF INDIVIDUAL CHLOROPLASTS**

At room temperature, the emission spectra of PSI and PSII show significant spectral overlap and cannot be distinguished using band-pass filtering as a straightforward approach. However, in a high-resolution fluorescence emission spectrum, the obtained spectral shape depends on the relative contributions of PSI and PSII, and hence the two subpopulations can be identified, compared, and distinguished from each other (**Figure 2A**). To survey the suitability of the SART approach for PSI/PSII intensity ratio determination, chloroplasts of immature and mature leaves of maize (*Zea mays*), a canonical C4 plant with Kranz anatomy consisting of mesophyll and bundle sheath cells, were used as model system. Maize chloroplasts of immature bundle sheath

**FIGURE 2 | PSI/PSII ratio in chloroplasts of living plant cells determined by SART. (A)** Average PSI/PSII ratio of chloroplasts in immature maize leaf tissue [left; calculated from 19 spectra recorded from mesophyll cells (MC) and 19 spectra recorded in bundle sheath cells (BSC), (*n* = 38, *p* = 0.1)] and in mature leaf tissue (right; calculated from 38 spectra recorded in MC and 59 spectra recorded in BSC). The upper left shows a brightfield image of a maize leaf section with MS and BSC cells. The upper right depicts a typical fluorescence intensity spectrum recorded from MC (x axis: emission wavelength [nm]). **(B)** Histogram of the calculated PSI/PSII intensity ratio of *A. thaliana* grown under long day (LD) and short day (SD) conditions. The histograms were calculated from 893 spectra in case of LD plants and 630 spectra in case of SD conditions. The histograms were fit with lognormal model functions with an adjusted *R*2-value of at least 0.95 (solid lines). **(C)** Average PSI/PSII intensity ratios in chloroplasts of *A. thaliana* wild type (Col0) and transgenic plants expressing maize plastidic NADP malic enzyme at different levels (MEm2 and MEm5 plants with 6 and 33 times increased NADP ME activity with regards to the wild type, respectively) grown under short day (SD) or long day (LD) conditions.

cells exhibit a high PSII activity (Lozier et al., 1971) while those of mature bundle sheath cells have established the characteristic agranal or only rudimentary grana morphology and show a low PSII activity (Woo et al., 1970). Using the *in vivo* SART approach we determined a low PSI/PSII ratio in bundle sheath cells of immature maize leaves, whereas this ratio rose substantially for bundle sheath cells of mature plants (**Figure 2A**). Based on this data, we validated that SART enables the determination of the PSI/PSII ratio in living plants. We therefore used SART to investigate the ability of the prototypical C3 plant *Arabidopsis thaliana* (Arabidopsis; *A. thaliana*) to adjust the PSI/PSII ratio to external and environmental cues.

To check the influence of the duration of the light period on the relative PSI/PSII luminescence, we grew *A.thaliana* ecotype Columbia-0 (Col0) under long day (LD) and short day (SD) conditions (16/8 and 8/16 h photoperiod, respectively). Spectra were recorded from plants at the same growth stadium in the second half of the light period. To prevent any adulteration of the results by spectra close to the noise or saturation limit of the spectrograph, we disregarded the most and least intense 5% of the recorded spectra for downstream processing. The calculated PSI/PSII ratios show a monomodal distribution that can be well explained by a lognormal model in both LD- and SD-grown plants (**Figure 2B**). As a *t*-test analysis might be misleading to identify whether the means of these distributions differ significantly, we performed a Z-score test using unbiased estimators (Zhou et al., 1997) to calculate the significance level for this lognormal distributions according to

with

$$\varphi\left(Z\right) = \frac{1}{\sqrt{2\pi}} \int\_{-\infty}^{Z} \exp\left(\frac{-\mathbf{x}^2}{2}\right).$$

−∞

*dx*

α = 2ϕ (*Z*); *Z* < 0

Plants grown under LD conditions show an overall significantly increased PSI/PSII ratio compared to plants grown under the more restrictive SD conditions. We calculated a Z-Score of *Z* = −12.57 translating into a significance level of 2.9∗10−36. These results indicate that *A. thaliana* plants adapt their photosynthesis apparatus, namely the apparent PSI/PSII ratio, depending on the available amount of light.

To investigate the capability of the *in vivo* adjustment to changes in internal cues, we compared the PSI/PSII ratio of *A. thaliana* wild type plants with that of two independent transgenic lines overexpressing the maize photosynthetic NADP-malic enzyme (MEm2 and MEm5 with 6 and 33 times higher NADP-ME activity Fahnenstich et al., 2007). As shown previously, the MEm plants present a sustained low level of the organic acids malate and fumarate, especially when grown in SD (Fahnenstich et al., 2007; Zell et al., 2010). As a consequence of the impairment in the supply of carbon in SD, the MEm plants show a dwarf growth phenotype, which is accompanied by more crowded thylakoid membrane systems and a lower PSII quantum yield compared to the wild type. The intensity of the changes described follows a dose response relationship with the level of NADP-ME activity (Zell et al., 2010). However, no differences in phenotype and all the above-mentioned parameters were found when the MEm plants were grown under LD conditions (Zell et al., 2010). For the determination of the PSI/PSII intensity ratio by SART, we grew the plants at both LD and SD conditions and recorded several hundred emission spectra per condition and plant. Again, the histograms of the calculated PSI/PSII intensity ratios obtained are monomodal and plausibly explained by a lognormal model (Supplemental Figure S1). Comparing the means of the PSI/PSII distributions, we found that MEm2 plants show no significant change in their PSI/PSII intensity ratio compared to wild type plants under LD conditions (significance level: 0.64). Furthermore, the MEm2 plants have a moderately enhanced PSI/PSII intensity ratio compared to wild type plants under SD conditions (significance level: 6.5∗10−8). This indicates that the slight overexpression of the plastidic NADP-malic enzyme has, if at all, only a moderate effect on the PSI/PSII intensity ratio in SD conditions. In contrast, the MEm5 plants show a pronounced drop in the PSI/PSII intensity ratio under LD conditions and slight increase under SD conditions (**Figure 2C**; significance level of 5.3∗10−27 and 5.8∗10−9 at LD and SD, respectively). This indicates that the chloroplasts of the MEm5 plants adapt to the carbon shortage by lowering the PSI/PSII intensity ratio under LD conditions. Since the quantum yield of PSII does not depend on the NADP-malic enzyme expression level when grown under LD conditions (Zell et al., 2010), this change in the PSI/PSII intensity ratio reflects an alteration in the relative PSI amount. Possibly, the alteration of the PSI/PSII intensity ratio in LD is one compensatory mechanism that enables the MEm5 plants to prevent a dwarfed growth phenotype.

## **USING FLUORESCENCE EXCITATION SPECTRO-MICROSCOPY (FExS) TO INVESTIGATE THE EFFICIENCY OF LIGHT HARVESTING COMPLEXES IN AN INDIVIDUAL CHLOROPLAST**

To investigate the individual efficiency of LHCs and their ability to adapt to changing environmental conditions, we applied FExS to 5-weeks-old *A. thaliana* wild type and MEm5 plants grown under SD conditions. The emission passband was chosen to match the PSII-emission. As shown in **Figure 3**, wild type and MEm5 plants showed three fluorescence excitation peaks between 520 nm and 620 nm (λpeak1 = 534 nm; λpeak2 = 568 nm; λpeak3 = 604 nm). We initially assumed these three peaks to originate in different pigments; however, there are no three pigments exhibiting their absorbance maxima in this wavelength regime. Rather, the peaks are assigned to the Qx(0, 0) and higher vibronic Qx and Qy transitions of the chlorophylls (Breton and Vermeglio, 1988; Fragata et al., 1988). The excitation spectra shown here reflect how much light is funneled to the PSII reaction centers upon absorbance at a given wavelength. Assuming that the LHCs and PSs form highly optimized functional units one expects that there is little variation in the composition of these cellular compounds and, as a consequence, this is expected to be reflected in a very constant excitation spectrum.

To establish if there is any adaptation of plants not only in the PIR but also inside the light harvesting apparatus, we recorded chloroplast excitation spectra *in planta* from wild type and from MEm5 plants. For data analysis, three Gaussians were fit to every spectrum as shown in **Figure 3** and the areas under the peaks were calculated in relation to the area under the first peak at 534 nm (peak S1).

As shown in **Figure 4A**, the spectra of wild type plants showed a rather broad distribution of the peak area ratios [2σ(S2:S1) = 0.73 ± 0.05, σ(S3:S1) = 1.60 ± 0.14, 2σ(S3:S2) = 0.96 ± 0.24). This indicates that light harvesting

highest transition energy (S1). Histograms of the peak ratio distribution of the excitation peak areas for wild type **(A)** and MEm5 plants **(B)**. Red: peak ratio S2:S1, green: peak ratio S3:S1; blue: peak ratio S3:S2. The histograms were fit by Gaussians (solid lines) and show normal distribution.

in living plants is by no means due to a rigid, uniform machinery but rather exhibits variations in terms of efficiency on a local basis. In contrast, for the MEm5 plants, the peak ratio distributions were significantly narrowed compared to the wild type [2σ(S2:S1) = 0.60 ± 0.01, σ(S3:S1) = 1.09 ± 0.08, 2σ(S3:S2) = 0.65 ± 0.04) (**Figure 4B**). Furthermore, the peak ratios exhibit an overall shift toward larger values for MEm5 plants compared to their wt counterparts. This means that overall, a higher fraction of fluorescence is excited at higher wavelength regions in MEm5 plants than in Col0.

## **DISCUSSION**

The results reported in this manuscript offer several insights into the adaptation potential of the photosynthetic apparatus of higher plants. First, our SART data substantiate previous findings that differences in the length of the light period in live plants lead to a reprogramming of the PSI/PSII ratio. We believe that the measured change in the PSI/PSII intensity ratio of LD- and SD-grown plants can be translated into a readjustment in the PSI/PSII ratio in plants because the quantum yield of Col0 plants grown in the applied light conditions does not change significantly (Zell et al., 2010). In contrast to previous studies, our data were obtained applying a non-invasive approach at ambient temperature conditions. As our data support the results of ultrastructural experiments, we consider our technique valid to monitor the PSI/PSII intensity ratio in living plant cells. We showed that SART can be used to investigate adaptation processes in chloroplasts of plants with changes in carbon homoeostasis with high sensitivity.

In LD conditions, MEm2 plants show no distinguishable growth and PSI/PSII intensity ratio phenotype from wild type plants, although they exhibit slightly lower levels of CO2 assimilation (Zell et al., 2010). Under this photoperiod, it appears that the photosynthetic productivity can still compensate for the physiological carbon depletion. Therefore, a significant adjustment of the PSI/PSII intensity ratio in MEm2 and wild type plants grown under LD conditions is not required. Contrarily, MEm5 plants, which exhibit a much higher level of NADP-ME, show clear changes in the PSI/PSII intensity ratio under LD conditions. Apparently, the carbon deficit under these conditions is already severe enough to provoke an effect in the composition of the photosynthetic apparatus. Interestingly, other parameters related to photosynthesis such as the total chlorophyll content and the PSII quantum yield are not different between MEm5 and wild type plants (Zell et al., 2010). Under SD conditions, there is a clear growth phenotype for both MEm2 and MEm5 plants. This is accompanied by a change in the PSI/PSII intensity ratio in MEm plants compared to wild type. MEm plants grown under SD conditions furthermore experience a decreased PSII quantum yield compared to wt plants which is progressive with the NADP-ME activity (Zell et al., 2010). Interestingly, the difference in the PSI/PSII intensity ratio of MEm2 and MEm5 plants is not linear with the NADP-ME activity. We observed that the PSI/PSII intensity ratio of MEm2 plants decreases from LD- to SD-grown plants whereas in MEm5 plants there is an increase. The altered PSII quantum yield might well affect the measured PSI/PSII intensity ratio. However, the fact that these two effects are not correlated is a strong indication that the observed change in the PSI/PSII intensity ratio originates in a relative amount of PSI/PSII. This finding is interpreted as follows: Under sufficient light conditions—as is the case in LD—there is enough energy supply to compensate for the carbon deficit allowing normal growth. In case of MEm2, there is no adaptation necessary due to the moderate overexpression of the NADP-ME. In MEm5, however, a detrimental growth phenotype may be avoided by physiological/molecular compensatory changes, being one of them the large reorganization of the photosynthetic apparatus. In SD conditions, the photosynthetic productivity of MEm chloroplasts can no longer overcome the reduced CO2 assimilation due to a large carbon starvation. Even a possible re-adjustment of the PSI/PSII ratio, which might reflect a general, adaptive response to limiting light conditions, cannot compensate for the carbon deficit.

A deeper insight into the LHCs of chloroplasts is obtained by application of the FExS approach, which allows identifying the direct observation of the amount of light that is funneled to the downstream excitation centers after excitation of a specific energy state. Our FExS data revealed that the energy transfer to the PSII reaction centers upon excitation in different wavelength regimes is not constant but exhibits significant variations within a plant. The three excitation peaks described in this study do not translate into the amounts of different pigments involved in light harvesting. Rather, we interpret the energy cascade in LHCs as complex multi-level entities. If the LHCs were uniformly assembled, no variations at all would be observed in the excitation peak ratios, just like in the excitation spectra of fluorophores in solution. The fact that we observe strong variations here means that the efficiency of downstream energy transfer upon excitation of a distinct energy level is not constant. As a consequence, a varying fraction of the collected light might migrate by pathways different than energy transfer to the PSII reaction center upon excitation, arguably by non-radiative deactivation of the excited state. The fluctuations observed here essentially reflect a different nanoenvironment of the excited pigments not observable in isolated systems. This indicates that the chlorophylls of the energy cascade involved in light harvesting are not rigidly embedded into the protein scaffold but rather represent a highly dynamic system *in vivo*. Interestingly, we found that MEm5 plants suffering from carbon starvation exhibit an overall altered photosynthetic efficiency in the probed spectral regime *in vivo* along with a narrowed distribution of the excitation peak ratios. This indicates an overall altered, more uniform composition of MEm5 chloroplasts compared to wild type plants. On microscopic level, wild-type plants seemingly maximize the light absorption and the subsequent energy transfer to the reaction centers. The MEm5 plants obviously present a limitation of this ability. Likely, the MEm5 plants are not able to dynamically re-organize their LHCs like the wild type due to the lack of sufficient carbon supply during the extended night period (Zell et al., 2010). This may reduce the capacity of the MEm5 plants to optimize the photosynthetic energy generation process. Together with the de-regulation of the PSI/PSII ratio, this could contribute to the carbon starvation phenotype of the MEm5 plants under SD conditions.

Taken together, our results suggest the potential for adaptations within the LHCs of land plants such the *A. thaliana* based on a different interaction network of the pigments involved in light harvesting, a mechanism differing from the chromatic adaptation previously reported to be due to alterations of the overall LHC pigment composition incyanobacteria (Kehoe and Gutu, 2006; Mascle-Allemand et al., 2010).

## **CONCLUDING REMARKS**

In this work, we presented two spectro-microscopic approaches, namely SART and FExS, to investigate for the first time photosynthetic adaptation processes in single chloroplasts of living plant cells in their native milieu at ambient temperature. Our data show that SART is a sensitive method for detecting and quantifying externally- (light conditions) and internally- (carbon homoeostasis changes) regulated adaptation processes of chloroplasts. This method is highly sensitive and is thus well suited to investigate the dynamics of PSI/PSII ratio of chloroplasts *in vivo*. Whereas SART spectroscopy enables the determination of the relative PSI/PSII amounts, FExS allows the disentanglement of the energy transfer processes in the LHCs. Both the PSI/PSII fluorescence intensity ratio and the energy transfer are demonstrated to be adjustable to external conditions such as light intensity and internal to cues, such as C4 acid deficiency, in a highly flexible manner.

## **ACKNOWLEDGMENT**

We acknowledge the support by the Deutsche Forschungsgemeinschaft (DFG) and the Open Access Publishing Fund of the University of Tübingen.

## **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www.frontiersin.org/journal/10.3389/fpls.2014.00292/ abstract

## **REFERENCES**

Allen, J. F., and Forsberg, J. (2001). Molecular recognition in thylakoid structure and function. *Trends Plant Sci.* 6, 317–326. doi: 10.1016/S1360-1385(01) 02010-6

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 02 July 2013; accepted: 04 June 2014; published online: 30 June 2014.*

*Citation: Peter S, Zell MB, Blum C, Stuhl A, Elgass K, Sackrow M, Subramaniam V, Meixner AJ, Harter K, Maurino VG and Schleifenbaum FE (2014) Photosynthesis in a different light: spectro-microscopy for in vivo characterization of chloroplasts. Front. Plant Sci. 5:292. doi: 10.3389/fpls.2014.00292*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Peter, Zell, Blum, Stuhl, Elgass, Sackrow, Subramaniam, Meixner, Harter, Maurino and Schleifenbaum. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Spectral analysis combined with advanced linear unmixing allows for histolocalization of phenolics in leaves of coffee trees

#### *Geneviève Conéjéro1, Michel Noirot 2 \*, Pascale Talamond3 and Jean-Luc Verdeil 1*

*1 Plant Cell Imaging platform PHIV UMR AGAP (Cirad, SupAgro, INRA), UMR B&PMP (INRA, CNRS, UM2, SupAgro), Montpellier, France*

*2 UMR PVBMT, (CIrad, IRD) La Réunion, France*

*3 Institut des Sciences de l'Evolution, UMR ISE-M (CNRS, IRD, UM2), Montpellier, France*

#### *Edited by:*

*Tobias Meckel, Technische Universität Darmstadt, Germany*

#### *Reviewed by:*

*Andreas Nebenführ, University of Tennessee, USA Hirokazu Tanaka, Osaka University, Japan*

#### *\*Correspondence:*

*Michel Noirot, Institut de Recherche pour le Développement, 911 Avenue Agropolis, 34394 Montpellier, France e-mail<EMAIL_ADDRESS>
An imaging method using spectral analysis combined with advanced linear unmixing was used to allow histolocalization of natural autofluorescent compounds such as hydroxycinnamic acid (chlorogenic acid) and xanthone (mangiferin) in living cells and tissues (mature coffee leaves). The tested method included three complementary steps: 1/ visualization of natural autofluorescence and spectrum acquisition with a multiphoton microscope; 2/ identification of some compounds using previous information on the chemical composition of the tissue, obtained from litterature; and 3/ localization of candidate compounds by spectral imaging. The second part of the study consisted of describing the histochemical structure of leaves during their development. This revealed very fast histochemical differentiation of leaves during the first week after their emergence. Lastly, young leaves of *Coffea pseudozanguebariae* (PSE), *C. eugenioides* (EUG), *C. arabica* (ARA) and *C. canephora* (CAN) were compared. This confirmed the presence of xanthone in PSE and EUG*,* but especially its precise tissue localization. This also highlighted the paternal CAN origin of the leaf structure in the allotetraploid species ARA*.* The limits and advantages of the method without staining are discussed relative to classical epifluorescence microscopy under UV light. This non-invasive optical technique does not require pretreatment and is an effective experimental tool to differentiate multiple naturally-occuring fluorochores in living tissues.

#### **Keywords: spectral analysis, multiphotonic microscope, autofluorescence, caffeoylquinic acids, xanthone,** *Coffea*

## **INTRODUCTION**

Plants produce a vast array of secondary metabolites such as phenolics that are estimated to comprise at least 8000 different chemicals (Jones et al., 2013). Many of them include low molecular weight phenolics, but also condensation products such as lignins and flavonoids. It is estimated that phenolics represent about 40% of all organic compounds circulating in the biosphere. These contribute to hardiness, color, taste, odor and many of them are of high economic value.

In the *Coffea* genus, caffeoylquinic acids (CQA) and dicaffeoylquinic acids (diCQA)-two secondary metabolites from the phenylpropanoid pathway-are of major economic importance in coffee production via two major species: *Coffea arabica* L. (ARA) (65–70% of the worldwide coffee production) and *C. canephora* Pierre ex Froehner (CAN). CQA and diCQA are indeed the most abundant soluble hydroxycinnamic acids (HQA) present in leaves and seeds (Clifford, 1985; Ky et al., 2001; Campa et al., 2012). Three isomers occur in each class according to the acylating residue positions, but the most important in terms of content is 5-*O-*caffeoylquinic acid, more commonly known as chlorogenic acid (Clifford, 1985) (see **Figure A** in supplementary material).

Another important xanthonoid phenol has been identified in leaves of *C. pseudozanguebariae* Bridson (PSE). This is *C*-glycosyl xanthone, or 2-*C*-β-D-glucopyranosyl–1,3,5,7 tetrahydroxyxanthen-9-one, more commonly known as mangiferin (Talamond et al., 2008, 2011; Campa et al., 2012) (see **Figure A** in supplementary material). As for chlorogenic acid (Bertrand et al., 2003; Mondolot et al., 2006), its content was found to decrease from young to mature leaves (Campa et al., 2012). Lastly, it has also been detected in other *Coffea* species such as *C. eugenioides* S. Moore (EUG) and ARA, but is absent in CAN leaves (Campa et al., 2012).

Great progress has been made in understanding the regulation of the expression of genes involved in phenol metabolism, but less is known about their spatial distribution at the tissular and cellular level. To date, phenol histolocalization is generally done using Neu's reagent, which binds with phenolics, emitting a specific greenish-white epifluorescence under UV light (Neu, 1956). As expected, the most intense greenish-white fluorescence is observed in juvenile coffee leaf blades (Mondolot et al., 2006; Campa et al., 2012), but its specificity toward HQA is low. By contrast, mangiferin histolocalization by epifluorescence does not require any reagents and has been directly observed through its autofluorescence (Talamond et al., 2011; Campa et al., 2012). As phenolics are autofluorescent compounds, it was suggested that microspectrometry allowing fluorescence spectra recording from ROI could be used to confirm their fluorescence signature (Hutzler et al., 1998). This can be achieved using either confocal (Hutzler et al., 1998) or two-photon microscopy (Talamond et al., 2011). The latter has the advantage of providing excellent three-dimensional spatial resolution of the location from which the fluorescence spectra can be obtained (Saadi et al., 1998). This imaging approach is not commonly used to localize phenolic compounds since fluorescence spectra obtained from plant samples are generated by the emission of many fluorophores with overlapping emission peaks. Fortunately, spectral unmixing methods using computer algorithms provide an opportunity to separate a spectral signal recorded from a single or a group of pixels containing a number of fluorophores into separate intensity signals of each of them.

In this paper, a new imaging approach was carried out to localize phenols *in situ* using a multiphoton microscope combined with spectral analysis and linear unmixing. This allows the differentiation of distinct fluorophores with highly overlapping emission spectra (Zimmermann et al., 2003; Garini et al., 2006). We developed such a method to localize 5-CQA and mangiferin, which are fluorescent compounds, in fresh mature coffee leaves. Nevertheless, it is important to choose plant material with a high proportion of 5-CQA in HQAs in order to minimize noise due to other HQAs. As the highest CQA/HQA ratio (93%) was found to characterize mature ARA leaves (estimated from Campa et al., 2012), we opted to use this material in the first part of the present study. The second part involved describing the differentiation of the histochemical structure of ARA leaves over time. Lastly, the histochemical structure of young leaves was compared between four *Coffea* species, i.e., PSE, EUG, ARA, and CAN. PSE was selected for its very low 5-CQA content (Bertrand et al., 2003) and high mangiferin content in leaves (Talamond et al., 2011; Campa et al., 2012). EUG and CAN were taken into account because they are putative parents of ARA (Lashermes et al., 1999). In addition, the four species constitutes a gradient for the chlorogenic acid content (Campa et al., 2012).

## **MATERIALS AND METHODS PLANT MATERIAL**

For *Coffea pseudozanguebariae* (PSE), *C. eugenioides* (EUG), *C. arabica* L (cv Bourbon, ARA) and *C. canephora* (CAN), leaves were sampled on coffee trees growing in the field at St Pierre, Réunion (France). Mature leaves were sampled on the third node from the branch tips, whereas young leaves were sampled at phase ϕ2 and ϕ3 (Lécolier et al., 2009).

## **HISTOLOGICAL AND IMAGING METHODS**

Leaf cross-sections (50μm) were obtained using an HM 650 V vibrating blade microtome (Microm, Waldorf, Germany) and then dipped in a glycerol/water (50:50) solution saturated with ascorbic acid to prevent oxidation. A multiphoton Zeiss 510 META NLO microscope (Zeiss, Iena, Germany), equipped with a laser Chameleon Ultra II Ti-Sapphire (Coherent, Santa Clara, California) and a 25x/0.8 Plan-Neofluar immersion objective, was used to obtain galleries of spectral images and emission spectra from fresh leaf sections. Spectral imaging was carried out with laser optimal excitation at 720 nm, reproducing laser UV monophotonic excitation in the 365–700 nm emission range. A set of 32 images was obtained, with each image being acquired with a separate narrow bandwidth (10.7 nm), representing the complete spectral distribution of the fluorescence signals for every point of the microscopic image. This procedure was performed on leaf cross-sections, as well as on pure chlorogenic acid (5-CQA) and mangiferin powders (Sigma-Aldrich, St Quentin Fallavier, France and Extrasynthese, Genay, France, respectively).

To check a possible impact of pH on spectral emission, some experiments using different pH (between pH = 5.5 and 10.5) were carried out in the case of the 5-CQA. There was no impact on the results (data not shown).

The spectral analysis was carried out using the advanced linear unmixing function (LSM 510 software) which separates mixed signals pixel by pixel using the entire emission spectrum of each defined autofluorescent compound in the sample. This function requires at least two spectra and was applied with the advanced iterative option and one residual channel. After spectral imaging acquisitions on leaf cross-sections, the advanced linear unmixing function allowed visualization with coded colors of the fluorescence of chlorogenic acid (5-CQA), mangiferin, and chlorophyll in cells based on their reference spectra.

Each experiment was repeated with 3–5 leaves per different stages or species.

## **RESULTS**

## **CHARACTERIZATION OF LEAF ANATOMY UNDER FLUORESCENCE**

The first step involved obtaining spectral images from mature leaf cross-sections of the ARA (*C. arabica* L cv Bourbon) (**Figure 1**). This image was the result of merging 32 images (channels) obtained between 365 and 700 nm and shows the whole fluorescence detectable in the leaf section without staining. The autofluorescence of cell walls, pigments and cuticle enabled visualization of the leaf anatomical structure.

**FIGURE 1 | Spectral image on leaf cross-section of the** *C. arabica* **cv. Bourbon obtained using a multiphoton microscope at 720 nm excitation.** AdE, adaxial epidermis; AbE, abaxial epidermis; C. cuticule; PP, palisade parenchyma; SP, spongy parenchyma; VB, vascular bundles. Scale bar = 50μm.

As described by Dedecca (1957), leaf cross-sections showed the classical foliar structure with the adaxial epidermis, palisade parenchyma, spongy parenchyma, and abaxial epidermis. Each epidermis was covered by a thin cuticle. Lastly, vascular bundles were also present on this cross-section.

The autofluorescence software coding function was then applied, thus generating the central part of **Figure 2**. Coding consisted of giving a wavelength-dependent color to each pixel with a proportional intensity to the pixel fluorescence intensity (Garini et al., 2006). This encoding clearly brought out the previously described leaf structure (**Figure 1**). Three groups of tissues were distinguished according to the autofluorescence intensity and color: 1/both epidermal tissues (adaxial and abaxial), which had low fluorescence; 2/parenchyma (palisade and spongy) was characterized by strong red fluorescence; and 3/vascular bundles showing strong blue fluorescence. Cuticles also showed high orange fluorescence. Lastly some small yellow fluorescent zones were observed within the palisade parenchyma.

The next step consisted of obtaining spectra from red, blue, yellow and orange zones (on the left and right parts of **Figure 2**). In the present case, each spectrum was characteristic of one pixel, symbolized by a cross on the figure, but it was also possible to define each emission spectrum from a small number of pixels, giving identical results. The next step involved looking for compounds that could explain the autofluorescence of these different ROI.

## **IDENTIFICATION AND LOCALIZATION OF 5-CQA, MANGIFERIN AND CHLOROPHYLL**

Reference spectral signatures were acquired on the microscope using controls of products known to be present in mature leaves, i.e., pure 5-CQA and mangiferin powders and chlorophyll extracts. **Figure 3** represents the reference spectra of 5-CQA, mangiferin and chlorophyll. Each molecule could be characterized by its absorption and emission spectra. Spectral signatures of 5-CQA, mangiferin, and chlorophyll were then stored in the Spectra Database. The emission spectrum of 5-CQA obtained with the laser system had the same profile and λmax emission (457 nm) as that obtained with a conventional spectofluorometer with a 300 nm excitation wavelength. Spectral signature of chlorophyll presented λmax at 670 nm, as expected. 5-CQA and mangiferin presented a wide spectral range with peaks at about 455 and 590 nm, respectively.

For the last step, the advanced linear unmixing process was carried out using chlorophyll, 5-CQA and mangiferin reference spectra (**Figure 4**). The composite image (V) was generated from four base images (I–IV), corresponding to the localization of chlorophyll (I), 5-CQA (II), mangiferin (III), and residual fluorescence (IV), respectively. Similarity is striking when comparing **Figures 2**, **4V**.

In fact, chlorophyll, 5-CQA and mangiferin spectra looked like ROI1, ROI2, and ROI3 of the **Figure 2**, respectively. Differences were nevertheless observed due to the fact that several autofluorescent compounds were present at a given pixel. For example, mangiferin spectra did not show a peak at 670 nm, suggesting that ROI3 resulted from both mangiferin and chlorophyll spectra.

High chlorophyll fluorescence was observed in chloroplasts of parenchyma cells (palisade and spongy), but not in the epidermis and leaf veins. 5-CQA fluorescence was mainly distributed in vascular bundles and in the cuticle of the abaxial epidermis, and to a lesser extent in some cells of the adaxial epidermis. It was not detected on the adaxial cuticle. Mangiferin fluorescence appeared in all parenchyma cells and in cuticles (adaxial and abaxial).

**FIGURE 2 | Coded autofluorescence in a** *C. arabica* **cv. Bourbon leaf blade cross-section observed using a multiphoton microscope at 720 nm excitation.** Four spectra emissions were recorded, corresponding to red (ROI1), blue (ROI2), yellow (ROI3) and orange (ROI4) colors, respectively on

chloroplasts, vascular bundle, unknow cellular structure and cuticle. AdE, adaxial epidermis; AbE, abaxial epidermis; C, cuticule; PP, palisade parenchyma; SP, spongy parenchyma; VB, vascular bundles. Scale bar = 50μm

## **HISTOCHEMICAL STRUCTURE VARIATIONS OVER A TIME COURSE IN ARA LEAVES**

In very young ARA leaves at the ϕ2 phase (Lécolier et al., 2009), the classical structure, as seen in **Figure 1**, was not achieved (**Figure 5I**) (see also **Figure BI** in supplementary material). Although the abaxial and adaxial epidermal layers were clearly visible, there was no clear differentiation between palisade and spongy parenchyma. Especially, the absence of clear differentiation characterized the histochemical structure of the leaves. All tissues showed the same blue color in coded autofluorescence (**Figure 5I**). Only the color intensity varied within leaves, thus highlighting the cell boundaries and vacuoles. Spectral analysis using chlorophyll, 5-CQA and mangiferin spectra showed the presence of chlorophyll at a very low level in the parenchyma (**Figure BI,** in supplementary material ). Moreover, the three spectra did not explain more than 50% of the fluorescence (intense residual fluorescence). Finally, two main differences characterized the histological distribution of 5-CQA and mangiferin: in the case of 5-CQA, the cell limits were not distinguishable, whereas vacuoles clearly appeared. By contrast, mangiferin was absent from cell walls thus highlighting these latter (**Figure BI** in supplementary material).

The histochemical structure changed very quickly in young leaves over a 4-day period (phase ϕ3), whereas leaves were always folded at the top of the branch, the classical histochemical structure with two parenchyma (palisade and spongy) became slightly visible (**Figure 5II**). Advanced linear unmixing confirmed the presence of chlorophyll in parenchyma, thus explaining the red zones observed in autofluorescence (see also **Figure BII** in supplementary material). The localisation of 5-CQA in both epidermis was also confirmed. Lastly, mangiferin was also present in vacuoles of both epidermal tissues, but at very low intensity. At maturity, the histochemical structure was definitively acquired, with large intercellular spaces in the spongy parenchyma (**Figure 5III**). The principal effect of age was the lowering of 5-CQA fluorescence in both epidermal layers (**Figure BI,** in supplementary material).

#### **HISTOCHEMICAL COMPARISON BETWEEN SOME COFFEA SPECIES**

Histochemical comparison using the advanced linear unmixing process concerned leaves of PSE, EUG, ARA "Bourbon" (ARA) and CAN. In mature leaves, the histochemical structure was similar in the four species (data not shown). This was not the case in young leaves (phase ϕ3). In this case, two groups of species could be defined, i.e., PSE and EUG vs ARA and CAN. In PSE (**Figure 6I**) and EUG (**Figure 6II**), there was strong histochemical differentiation between adaxial and abaxial epidermal tissues at the vacuole level, based on 5-CQA and mangiferin, respectively (**Figure 6I**). Nevertheless, mangiferin was also present in the spongy parenchyma and, at a lower level, in palisade parenchyma. See also splitted image of PSE leaf in **Figure BII** (supplementary material). In contrast, ARA (**Figure 6III**) and CAN (**Figure 6IV**) did not show mangiferin in adaxial and abaxial epidermal tissues and 5-CQA was strongly present in vacuoles of both of these tissues. In fact, mangiferin was only observable in parenchyma, but as small vesicles.

## **DISCUSSION**

#### **ADVANTAGES AND LIMITS OF THIS NEW IMAGING APPROACH**

Spectral analysis, combined with the advanced linear unmixing, provides an efficient tool to localize UV-fluorescent metabolites in living plant tissues. Based on the biochemical results obtained on total leaf extracts, i.e., the strong presence of 5-CQA and

**FIGURE 4 | Spectral analysis by linear unmixing method using chlorophyll, 5-CQA and mangiferin spectra.** Merged image V was splitted into four base images (I, II, III, and IV). Base image I showed histolocalization of chlorophyll, whereas base images (II) and (III) represented histolocalization of 5-CQA and mangiferin, respectively. Finally, base image (IV) depicted other fluorescent compounds (residual fluorescence). AdE, adaxial epidermis; AbE, abaxial epidermis; Scale bar = 50μm.

**FIGURE 5 | Histochemical structure of ARA (***C.arabica* **cv. Bourbon) leaves at very young (I), young (II), and mature (III) stages.** All these spectral images showed autofluorescence obtained using a multiphoton microscope at 720 nm excitation. AdE, adaxial epidermis; AbE, abaxial epidermis. Scale bar = 50μm.

mangiferin (Campa et al., 2012), the imaging method allowed their spatial localization in mature and young leaves of several *Coffea* species.

The main advantage of this histological approach was the possibility of reaching intact internal tissues within the cross-section. Images were acquired from selected nascent intact cells and tissues far from superficial layers containing damaged and oxidized cells due to cutting. Consequently, and contrary to standard methods using the Neu's reagent, the superficial oxidation of phenols had no effect on the images. The second advantage was the possibility of localizing, at the tissular and cellular level, a particular chemical species characterized by its own emission spectrum. For example, Neu's reagent cannot discriminate between the different types of phenol, whereas the imaging method permitted specific localization of 5-CQA. Moreover, the method does not require any previous extraction of chlorophyll or other pigments. Especially, as mentioned by Zimmermann et al. (2003) and Berg (2004), spectral imaging, combined with advanced linear unmixing, is a powerful method to simultaneously localize several metabolites showing overlapped spectra.

Nevertheless, some caution is required in the interpretation. As any powerful method, the use of spectral imaging combined with linear unmixing has some limits that must be taken in consideration for image interpretation. For example, if metabolites such as isomers or dimers have similar UV-fluorescent spectra, their specific localization is not possible. This was the case for diCQA (dimer of 5-CQA) that had the same spectral signature than 5-CQA. In such situation results coming from biochemical

analysis must be considered to support image interpretation. In our study, it was only because quantification by HPLC showed that 5-CQA represented more than 80% of total HQA in mature ARA whose blue zones shown in **Figure 4II** could mainly reflect the 5-CQA localization.

In fact, the present localization of the 5-CQA by spectral analysis could concern both CQA and diCQA which in young leaves constitutes most of HQA. This problem is also true for classical histochemical analysis using Neu's reagent. Nevertheless, the advantages of spectral analysis, as discussed above, were still relevant. Lastly, the absence of HQA or mangiferin in a tissue or cell was an important result. Indeed, spectral imaging allows detection of very low content. This was the case when visualizing 3-CQA and 4-CQA in Sigma-Aldrich powder of 5-CQA, but also using a dilution gradient (data not shown).

## **PRACTICAL APPLICATION OF SPECTRAL ANALYSIS TO REVEAL THE HISTOCHEMICAL LEAF STRUCTURE AND FOR BETWEEN SPECIES COMPARISONS**

The very rapid change in the histochemical leaf structure was the first novel result of the study. In less than 3 days, leaves developed from phase ϕ2 to phase ϕ3 (Lécolier et al., 2009) and their histochemical structure changed substantially. Beyond phase ϕ3, the histological structure was slightly modified and this concerned the spongy parenchyma. In parallel, the blue intensity decreased markedly in both epidermal tissues. This evolution from phase ϕ3 to leaf maturity has already been observed in CAN using HPLC (Mondolot et al., 2006). In young leaves (phase ϕ3), 5-CQA and diCQA contents were found to be 2.94 and 2.75% dry matter basis (dmb), respectively, whereas they were 0.58% and 0.14% dmb in mature leaves, respectively.

The differences in 5-CQA and mangiferin histolocalizations between species represented the second novel result of this study. The strong presence of mangiferin in young leaves of PSE was confirmed (Talamond et al., 2008, 2011; Campa et al., 2012). In these leaves (phase ϕ3), the mangiferin content was 8.6% dmb and decreased up to 5.6% dmb in mature leaves (Campa et al., 2012). Mangiferin has also been localized using UV epifluorescence microscopy (Talamond et al., 2011; Campa et al., 2012). The present results showed a more precise histolocalization of mangiferin in small vesicles of the parenchyma and in vacuoles of the abaxial epidermis, but not in the adaxial epidermis. Conversely, 5-CQA was only localized in the abaxial epidermis. 5-CQA was most abundant and constituted an important reservoir of phenols but its role has not yet been clearly defined. This compound could have at least two different functions, being involved in lignin synthesis, producing the precursor of the lignin structure (guaiacyl and syringyl units), while also being a protective agent against UV or aggressors (Hoffmann et al., 2003). Further experiments are under way to substantiate these assumptions.

The second important result was obtained through the species comparison, highlighting similarities between PSE and EUG, and between CAN and ARA. As EUG and CAN are maternal and paternal parents of ARA (Lashermes et al., 1999), respectively, our work clearly highlighted the paternal origin of the histochemical structure of ARA leaves.

## **ACKNOWLEDGMENTS**

This study was financially supported by the European Union, the Conseil Régional of Réunion and the French Institut pour la Recherche et le Développement (IRD).

## **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www.frontiersin.org/journal/10.3389/fpls. 2014.00039/abstract

#### **Figure A | Chemical structures of 5-CQA and mangiferin.**

**Figure B | (I)** Spectral analysis by linear unmixing method using 5-CQA, mangiferin and chlorophyll spectra on ARA (*C. Arabica* "Bourbon") very young leaf splitted image. **(II)** Spectral analysis by linear unmixing method using 5-CQA, mangiferin and chlorophyll on PSE (*C. pseudozanguebariae*) leaf: splitted image.

## **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 29 November 2013; accepted: 28 January 2014; published online: 18 February 2014.*

*Citation: Conéjéro G, Noirot M, Talamond P and Verdeil J-L (2014) Spectral analysis combined with advanced linear unmixing allows for histolocalization of phenolics in leaves of coffee trees. Front. Plant Sci. 5:39. doi: 10.3389/fpls.2014.00039*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Conéjéro, Noirot, Talamond and Verdeil. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## An update: improvements in imaging perfluorocarbon-mounted plant leaves with implications for studies of plant pathology, physiology, development and cell biology

*George R. Littlejohn1\*, Jessica C. Mansfield2 , Jacqueline T. Christmas 3 , EleanorWitterick1, Mark D. Fricker 4 , Murray R. Grant 1, Nicholas Smirnoff 1, Richard M. Everson3 , Julian Moger 2 and John Love1\**

*1 Division of Plant and Microbial Sciences, School of Biosciences, University of Exeter, Exeter, UK*

*2 School of Physics, University of Exeter, Exeter, UK*

*3 Computer Science, University of Exeter, Exeter, UK*

*4 Department of Plant Sciences, University of Oxford, Oxford, UK*

#### *Edited by:*

*Markus Schwarzländer, University of Bonn, Germany*

#### *Reviewed by:*

*Viktor Zarsky, Charles University, Czech Republic Jin-Long Qiu, Institute of Microbiology – Chinese Academy of Sciences, China David C. Logan, Université d'Angers, France*

#### *\*Correspondence:*

*George R. Littlejohn and John Love, Division of Plant and Microbial Sciences, School of Biosciences, University of Exeter, Geoffrey Pope Building, Stocker Road, Exeter EX4 4QD, UK*

*e-mail<EMAIL_ADDRESS><EMAIL_ADDRESS>
Plant leaves are optically complex, which makes them difficult to image by light microscopy. Careful sample preparation is therefore required to enable researchers to maximize the information gained from advances in fluorescent protein labeling, cell dyes and innovations in microscope technologies and techniques. We have previously shown that mounting leaves in the non-toxic, non-fluorescent perfluorocarbon (PFC), perfluorodecalin (PFD) enhances the optical properties of the leaf with minimal impact on physiology. Here, we assess the use of the PFCs, PFD, and perfluoroperhydrophenanthrene (PP11) for *in vivo* plant leaf imaging using four advanced modes of microscopy: laser scanning confocal microscopy (LSCM), two-photon fluorescence microscopy, second harmonic generation microscopy, and stimulated Raman scattering (SRS) microscopy. For every mode of imaging tested, we observed an improved signal when leaves were mounted in PFD or in PP11, compared to mounting the samples in water. Using an image analysis technique based on autocorrelation to quantitatively assess LSCM image deterioration with depth, we show that PP11 outperformed PFD as a mounting medium by enabling the acquisition of clearer images deeper into the tissue. In addition, we show that SRS microscopy can be used to image PFCs directly in the mesophyll and thereby easily delimit the "negative space" within a leaf, which may have important implications for studies of leaf development. Direct comparison of on and off resonance SRS micrographs show that PFCs do not to form intracellular aggregates in live plants.We conclude that the application of PFCs as mounting media substantially increases advanced microscopy image quality of living mesophyll and leaf vascular bundle cells.

**Keywords: perfluorocarbon,** *Arabidopsis***, multi-photon, confocal, microscopy, imaging, perfluoroperhydrophenanthrene**

## **INTRODUCTION**

Advances in microscopy have made *in vivo* biological imaging increasingly important in recent years. A diverse palette of chemical labels and genetically encoded fluorescent reporters and biosensors, coupled with advanced microscopy techniques, including laser scanning confocal microscopy (LSCM), twophoton fluorescence (TPF) microscopy and label-free imaging techniques such as second harmonic generation (SHG) microscopy and stimulated Raman scattering (SRS) microscopy have enabled unprecedented analysis of living cell dynamics (Shaw and Ehrhardt, 2013). Despite these technical advances, tissues that are rich in airspaces, such as those of plant leaves or animal lungs, remain difficult to image because of the optical aberrations that result from the complex structure of such tissues. As these tissues are typically sites of active metabolism and often targets

for pathogens, it is essential to develop relatively simple *in vivo* methods to circumvent these imaging problems.

The spongy mesophyll of higher plant leaves is located adjacent to the lower epidermis, contains numerous airspaces and may be several cell layers thick (**Figure 1A**). These characteristics result in light refraction within the mesophyll and a progressive attenuation of light transmission through the tissue, producing optical aberrations that impair confocal image quality (Feijó and Moreno, 2004; Inoue, 2006; Cheng, 2006). When examining fixed leaves these aberrations are minimized because the fixatives infiltrate the tissue and minimize the optical phase transitions within. Previously, we have shown that it is possible to infiltrate living leaves with perfluorodecalin (PFD, **Figure 1B**) and thereby significantly improve the resolution of LCSM images of the mesophyll while affecting only minimally cellular physiology (Littlejohn et al., 2010). We

have had numerous positive reports from users of PFD and some studies have now been published (Johnson et al.,2011;Knapp et al., 2012; Tschiersch et al., 2012; Carrión et al., 2013; Gest et al., 2013; Hoepflinger et al., 2013; Hutt et al., 2013; Mansfield et al., 2013; Wright et al., 2013).

adaxial epidermis; st, stomatal pore; sp, spongy mesophyll; a.s,

In this paper, we report that the optical qualities of plant mesophyll can be further improved by infiltration with perfluoroperhydrophenanthrene (PP11, **Figure 1C**), a perfluorocarbon (PFC) that has a refractive index that is better matched to that of living cells. To measure the improvement in image quality more objectively, we developed a method of autocorrelation that quantifies the sharpness of the images acquired at varying depths within the leaf. Finally, using SRS microscopy we have shown that both PFD and PP11 are undetectable in living cells, but infiltrate the mesophyll airspaces homogenously. Furthermore, SRS imaging of PFCs *in vivo* may be used to delimit the "negative space" within plant leaves, i.e., the area within the leaf that is involved in gaseous exchange and pathogen invasion. We have advocated the application of PFC mounting of samples to studies of pathogenesis in plants and experiments to date are promising. For example, **Figure 1D** shows an example image of an *Arabidopsis thaliana* leaf infected with the pathogenic bacterium, *Pseudomonas syringae* pv. tomato (DC3000 expressing GFP), taken under the same conditions as those used in Hutt et al. (2013).

## **MATERIALS AND METHODS**

#### **PLANT CULTURE AND SAMPLE MOUNTING**

*Arabidopsis thaliana* (Col-0 ecotype) and transformants that stably and constitutively express a cytoplasmically localized "Venus" yellow fluorescent protein (SEYFP-F46L; Nagai et al., 2002) were used in this study. Seeds were surface sterilized for 3 min with 70% ethanol and then for 5 min with 10% sodium hypochlorite. Seeds were washed five times in water and suspended in 0.1%

agar. Seeds were stratified at 4◦C, in the dark, for 48 h before being sown on compost and grown at 20◦C, in a 16 h/8 h light/dark photoperiod.

Mature leaves were excised from plants aged approximately 3-weeks and sections floated in H2O, PFD, or PP11 for 5 min according to the methods described in Littlejohn et al. (2010), Littlejohn and Love (2012). Samples were mounted in the same medium and imaged by LSCM, TPF, SHG, and SRS microscopy.

#### **LSCM IMAGING**

Scale bar is 25 μm.

Confocal imaging was performed using a Zeiss Axiovert 510 Meta LSCM equipped with a 40x/1.30 oil DIC immersion C-Apochromat lens. Immersion medium was Zeiss immersol. Light paths and wavelengths were controlled by a 458/514 nm dichroic mirror. The pinhole was set at 70 μm. Images were integrated and processed using Zeiss 510 software. Images of Venus and chlorophyll fluorescence in intact *Arabidopsis* leaves were collected with excitation at 514 nm using a 30 mW argon laser, 6.1 A, 21.8% transmission intensity. Emission was recorded at 518–604 nm for Venus and at 647–690 nm for chlorophyll. Z-stacks containing 100 z-planes taken with 1 μm step size were collected for each of five samples incubated in PFD, PP11, or H2O for 5 min before imaging and mounted in the same medium for imaging. Figures were assembled in Adobe Indesign. **Figure 2** LSCM images were generated by using the "cut" function in Zeiss LSM Image Browser software, where the plane presented represents a cut through the entire z-stack from top to bottom taken at an angle of −24◦. The images therefore show a progression through the stack from top to bottom. Single z-planes are presented in **Figure 3**. GFP-expressing *Pseudomonas* (made according to Lambertsen et al., 2004) and chlorophyll shown in **Figure 1D** were excited respectively with 488 and 633 nm laser lines and emission captured from 505 to 570 nm (GFP)

taken of samples mounted in H2O, PFD and PP11 respectively. Images were generated by using the "cut" function in Zeiss LSM Image Browser software, where the plane presented represents a cut through the entire z-stack from top to bottom taken at an angle of −24◦. The images therefore show a

chlorophyll, shown in red. **(D–F)** 2-photon micrographs taken of samples mounted in H2O, PFD, and PP11 respectively. **(G–I)** SHG micrographs taken of samples mounted in H2O, PFD, and PP11 respectively. **(D–I)** are presented as projections of z-stacks. Scale bars are 50 μm.

and 647–711 nm (chlorophyll). The GFP-*Pseudomonas* Z-stack projected in **Figure 1D** was 30 μm deep, with a 0.25 μm step size and was captured with a C-Apochromat 63x/1.2W Corr M27 lens. The projection was made in Zeiss LSM510 software.

#### **RAMAN SPECTROSCOPY**

Prior to SRS imaging the Raman spectra of the PFCs were obtained using a Renishaw RM100 Raman microscope (Renishaw plc, UK), with a 785 nm diode laser and 1200 line/mm spectral grating, giving a spectral resolution of 1 cm−1.

#### **STIMULATED RAMAN SCATTERING MICROSCOPY**

Stimulated Raman scattering microscopy required two, pulsed laser beams; one at a longer wavelength (Stokes beam) and the other at a shorter wavelength (pump beam). The difference between the wavelengths of the pulsed lasers was tuned to correspond to the energy of a Raman vibration of interest. In our system a 1064 nm pico-second laser (PicoTrain HighQ laser) provided the Stokes beam and the output from an optical parametrical oscillator OPO (Levante Emerald APE) – pumped by the frequency doubled output of the picoTrain laser – provided the pump beam. The pump beam was tuned to a wavelength of 991.4 nm, which excited the C–F vibrations at 687.5 cm−1.

The amplitude of the Stokes beam was modulated at 1.7 MHz using an EOM. When SRS occurred within the sample, a modulated decrease in pump beam intensity [stimulated Raman loss; Moger et al. (2012)] was detected.

A modified laser scanning confocal microscope (Fluoview 300 IX71 Olympus) was used for imaging the SRS. The objective was a 60x 1.2 NA water immersion objective (UPlanS Apo Olympus). The transmitted light from the sample was collected with a 60x 1.0 NA water-dipping condenser and detected using a photo-diode

**(A–C)** typical autocorrelation results for single samples mounted in H2O, PFD, and PP11 respectively and imaged as z-stacks 100 μm deep. **(D–F)** representative images taken at a depth of 25 μm from samples mounted in H2O, PFD, and PP11 respectively **(G–I)** representative images taken at a depth of 50 μm from samples mounted in H2O, PFD, and PP11 respectively. Signal was collected from 518 to 604 nm for Venus, and is displayed in green and at 647–690 nm for chlorophyll, shown in red. **(J–L)** representative images taken at a depth of 75 μm from samples mounted in H2O, PFD, and PP11 respectively. **(M–O)** range and median plotted for all samples (*n*=5) mounted in H2O, PFD, or PP11 for red, green, and total signal respectively. Scale bars are 50 μm.

with a 70 V reverse bias (FDS1010 thorlabs). The 1064 nm Stokes beam was blocked from reaching the photo-diode using a band pass filter coherent anti-Raman scattering (CARS; 890/220 nm, Chroma technologies). A lockin amplifier was used to separate out the modulated SRL signal and the output from this was connected to the computer and imaging software. Raman Image stacks were acquired at a 512 × 512 pixel resolution and a 256 μm × 256 μm scan area with 1 μm separation between optical slices.

## **TWO-PHOTON FLUORESCENCE AND SECOND HARMONIC GENERATION MICROSCOPY**

Two-photon fluorescence and SHG were performed using the customized microscope described above. Excitation was provided by mode-locked femtosecond Ti:sapphire oscillator (Mira 900D; Coherent, USA) which produced 100-fs pulses at 76 MHz. The central wavelength of the fs beam was 800 nm with an average power at the sample of 5–30 mW. TPF and SHG were spectrally separated from the 800 nm excitation beam by a dichroic mirror (670dcxr; Chroma Technologies). After this, different bandpass filters were used to enable either TPF signal (CG-BG-39-1.00-1 and F70-500-3-PFU; CVI Melles Griot, UK) or SHG signal (F10-400- 5-QBL; CVI Melles Griot, UK) to reach the Hamamatsu R3896 photomultiplier tube.

## **QUANTIFICATION OF IMAGE CLARITY IN THE Z-AXIS**

The clarity of each image in relation to its position in the z-axis (i.e., into the tissue) was quantified using textural analysis. For each image, the "green" and "red" color channels were analyzed separately and in combination. A gray-scale spatial dependence matrix (Haralick et al., 1973), labeled "M," was constructed from the intensities (ranging from 0 to 255) of each pixel that composed each image and for each channel. The element M*r*,*c*, at row *r* and column *c* of this matrix is a count of how many times a pixel with intensity *r* has a pixel of intensity *c* in its immediate neighborhood, defined as the 8 pixels surrounding the measured pixel. Normalizing this matrix resulted in a set of joint probabilities of pixels with intensities *r* and *c* within the neighborhood. The correlation value associated with this normalized matrix (see Haralick et al., 1973) gives a measure of how closely correlated the intensity of a given pixel is with those of its neighboring pixels. The correlation measure was calculated for each image in acquired z-stacks, 100 μm deep, with a 1 μm z-resolution for leaves mounted in H2O, PP11 and PFD stacks, and separately for the red, green, and combined red/green channels.

## **RESULTS**

## **PERFLUOROCARBON MOUNTING MEDIA IMPROVE THE OPTICAL RESOLUTION OF DIFFERENT MODES OF LASER SCANNING MICROSCOPY**

We compared the image resolution of micrographic z-stacks acquired using LSCM, TPF, and SHG microscopy (**Figure 2**). TPF and SHG are non-linear optical techniques, which involve the simultaneous absorption of two or more photons. All these techniques are intrinsically confocal and generally use infra-red lasers instead of visible or UV lasers to excite fluorophores (Nandakumar et al., 2009). These techniques are considered advantageous compared to single-excitation confocal microscopy because

they can deliver improved imaging depths within scattering tissues and reduced photodamage within samples.

Two-photon fluorescence is similar to single photon fluorescence, but it requires the simultaneous absorption of two near infra-red photons, rather than a single UV or visible spectrum photon, to generate a fluorescent signal (Diaspro and Sheppard, 2002).

Second harmonic generation involves the simultaneous absorption of two near infra-red photons and the emission of a single visible photon with half the wavelength of the infra-red photons. This process only occurs in structures which lack inversion symmetry. In plant tissues these structures include cellulose and starch grains (Mizutani et al., 2000; Brown et al., 2003; Cox et al., 2005).

Under all modes of microscopy tested, PP11 and PFD outperformed H2O as an *in vivo* mounting medium, by infiltrating the mesophyll airspaces and smoothing the optical transitions within the mounted leaves. Moreover, we noted that PP11 outperformed PFD, which we ascribe to the refractive index of PP11 being closer to that of living cells compared to PFD. We were routinely able to acquire images from a depth of 100–135 μm within the leaf by LSCM and two photon imaging using PP11 as a mounting medium, which is greater than half the thickness of a leaf of a 3 week-old *Arabidopsis* plant and tests in with rice leaves allow imaging through the entire thickness of the leaf. The use of PFCs as a mounting medium also allowed the acquisition of SHG signals from chloroplast starch, which was not possible for samples mounted in H2O. This may, in itself, represent an important technique for the study of starch in leaves and statoliths in root cells.

To quantify the apparent advantage of using PFC mounting media to image within the mesophyll, we performed autocorrelation analysis on LSCM micrographs acquired at varying depths in *Arabidopsis* leaves. In this case, the fluorescence emission for both cytoplasmically localized Venus (Nagai et al., 2002) and chlorophyll were recorded. The pixel-by-pixel autocorrelation enabled an objective quantification of image quality. Using this method, we noted a wavelength-dependent improvement in resolution when samples were mounted in PFCs, compared to H2O (**Figure 3**). Images recorded deeper in the sample are more greatly affected by noise, which tends to be uncorrelated, and hence the correlation measure is low compared with the crisper images recorded closer to the surface. Median values for the autocorrelation demonstrate that PP11 and PFD outperform H2O and PP11 performs better than PFD, with a greater benefit seen in imaging Venus, compared with chlorophyll (**Figures 3M–O**). Most interestingly, the use of PFCs as mounting medium reduced the range of autocorrelation values obtained, showing that the images acquired when samples are mounted in PFD or in PP11 are not only clearer, but more consistent between replicate samples.

Normal cytoplasmic streaming and chloroplast movement was observed in all the experiments performed in this investigation, which is consistent with previous observations that the use of PFCs has a minimal effect on leaf physiology compared to mounting leaf samples in H2O (Littlejohn et al., 2010). This is also evidenced by the differences in chloroplast position seen between z-stacks represented in **Figures 4G–J**.

## **STIMULATED RAMAN SCATTERING IMAGING OF PERFLUOROCARBON DISTRIBUTION**

TPF and SHG and SRS are non-linear optical techniques, which involve the simultaneous absorption of two or more photons. All these techniques are intrinsically confocal and generally use infrared lasers instead of visible or UV lasers to excite fluorophores.

Stimulated Raman scattering is a chemical-specific technique which relies on stimulating Raman active molecular bond vibrations. SRS requires stimulation by two laser beams at different wavelengths (pump and Stokes beams), with the difference in wavelength between the two beams set to correspond to the energy of the molecular bond vibration of interest. When this condition is met, SRS occurs and results in a loss of intensity in the higher energy pump beam (stimulated Raman loss) and an equal increase in intensity in the Stokes beam (stimulated Raman gain). This process is detected by modulating one of the beams and detecting the modulations in intensity in the second beam using a lockin amplifier (Freudiger et al., 2008).

Perfluorocarbons readily infiltrate leaf airspaces. To ascertain whether PFCs remain localized in the airspaces of the mesophyll or are capable of also penetrating living cells, we imaged the PFC distribution in infiltrated *Arabidopsis* leaves using SRS. Prior to imaging, we demonstrated that Raman spectra may be used to identify specific peaks, which allow the C–F bonds in the PFC and the C–H bonds found in *Arabidopsis* surface waxes, in cellulose and in β-1-3-glucan to be easily distinguished (**Figures 4A–D**). PFC mounting media are therefore compatible with Ramanbased imaging of biological molecules (**Figures 4E,F**). Images of the "ON" and "OFF resonance confirmed that PFCs was homogeneously distributed in the airspaces between mesophyll cells (Littlejohn et al., 2010). However, the sensitivity of this technique enables us also to confirm that PFCs do not appear to form intracellular aggregates, and are therefore unlikely to penetrate beyond the apoplast into the protoplasm (**Figures 4G–J**). From a structural perspective, SRS imaging of PFC distribution in leaves has the added advantage of delimiting the mesophyll airspaces, potentially enabling a more intricate understanding and modeling of gas flow dynamics in leaves. The chloroplasts appear in both the on and off resonance SRS images due to their strong two-photon absorption (TPA). In this process they are absorbing one photon originating from the pump beam (either 991 or 989 nm on or off resonance) and one photon

**FIGURE 4 | Stimulated Raman scattering imaging of perfluorocarbons** *in vivo***. (A–D)** Raman spectra taken at 785 nm, comparing spectra of PFD and **(A)** PP11, **(B)** Beta 1–3 glucan, **(C)** cellulose, and **(D)** hexane extracted *Arabidopsis* leaf wax. Peaks labeled one and two were used to tune imaging to C–F and C–H bonds respectively. **(E,F)** SRS images representing

Z-stacks taken through *Arabidopsis* leaves mounted in **(E)** PFD and **(F)** PP11. **(G,H)** show respectively, off and on resonance images of an *Arabidopsis* leaf mounted in PP11. **(I,J)** show respectively, off and on resonance images of an *Arabidopsis* leaf mounted in PFD. Scale bars are 50 μm. Asterisks denote airspaces.

from the stokes beam (1064 nm). As the absorption spectrum of the chloroplasts is so broad, tuning the wavelength of the excitation laser does not affect the strength of the chloroplast signal.

Some of the chloroplasts may appear to be on the extracellular side of the red lines (cell walls) delineating the cells in the images.

This is due to the strong out-of-focus signal from chloroplasts; normally 2-photon excitation is constricted to a small, defined volume and therefore no pinhole is used to filter out-of-focus information. However, as explained, the absorption of the chloroplasts is so strong that where the chloroplast signal overlaps with that of the PFC, this is most likely an imaging artifact due to the strength of the TPA signal from the chloroplasts compared to the red fluorescence in the cells and the SRS from PFC, particularly as the phenomenon is seen in both on and off images. As the Chloroplast signal is very intense signal from adjacent image frames could be leaked into selected image frame (resolution in z direction is slightly worse than 1 um which is the image step size here). This has the effect of making the chloroplasts appear to be the "wrong" side of the red cell walls.

We cannot categorically exclude the possibility that PFC have penetrated the cells, but consider it to be unlikely as otherwise the TPS from the PFC would be much more ubiquitously distributed.

## **DISCUSSION**

In this investigation, we show that mounting living leaves in the PFCs, PFD, and perfluoroperhydrophenanthrene (PP11) quantifiably improves the clarity and consistency of images acquired from the mesophyll, for a number of laser-based microscopy techniques including LSCM, 2-photon microscopy, SHG, and SRS.

PFD and PP11 are non-toxic and possess a great carrying capacity for O2 and CO2. PFCs are not miscible with aqueous solutions, which are a disadvantage when trying to deliver bioactive compounds to cells in the leaf, however, this does suggest that PFCs may be of great use as they will not dilute metabolites or signaling molecules present in the cell wall, which could be easily perturbed by aqueous mounting media.

Both PFCs used in this investigation have low surface tensions (19.3 dyne cm−1 for PFD and 21.6 dyne cm−1 for PP11 compared to 72.8 dyne cm−1 for H2O; Sargent and Seffl, 1970) that are lower than the 25–30 dyne cm−1 required to passively overcome the stomatal barrier (Schönherr and Bukovac, 1972) and readily infiltrate the apoplastic space. This infiltration smooths the optical phase transitions within the mesophyll, resulting in reduced noise and quantifiably clearer images. Moreover, the properties that PFD has displayed for mesophyll – easy infiltration into the tissue, significant improvement in *z*-plane resolution and nontoxicity – may be exploited for more general, *in vivo* imaging of air-filled or heavily vascularised animal tissues, such as insect spiracles or vertebrate lung, where gaseous exchange is also important and which are primary target for microbial infection. Similarly, PFD may be used advantageously for the perfusion and imaging within organ cavities.

The improvement in image clarity obtained by mounting samples in PP11 is greater than for PFD, which we ascribe to a closer matching of the refractive index of PP11 (1.334) with that of leaf tissue, which has been estimated as 1.4–1.6 depending on wavelength of incident light (Paillotin et al.,1998; Feret et al.,2008) compared to that of PFD (1.313). This was shown by autocorrelation analysis of pixel intensities in LSCM images. This analysis also demonstrated that the use of PP11 is particularly advantageous for imaging shorter wavelengths (i.e., the "green" channel; **Figure 3**). Increased fluorescence transmission of shorter wavelengths may have important advantages when more than one fluorophore is imaged simultaneously, for example in analyses that require Forster resonance energy transfer (FRET) or co-localisation.

We anticipate that the general improvement in image quality conferred by mounting in PFC media could be further enhanced through an even closer match between the refractive indices of living cells and the PFC mounting medium. 2,2 -thiodiethanol (TDE) has previously been used in thisfashion (Staudt et al.,2007); in aqueous solution at varying concentrations, TDE provides a suite of colorless mounting media with tuned refractive indices from 1.33 to 1.52. Although TDE is not compatible with living samples and used preferentially with fixed specimens, it raises the possibility of a tuneable mounting system composed of two PFCs, each with refractive indices that bracket those of living specimens. These complementary PFCs may then be combined in set proportions to make a mounting medium in which the refractive index is matched to that of any sample.

The presence of C–F bonds in PFCs can be readily distinguished from the C–H bonds found in biological molecules. Consequently, PFCs does not impair the imaging of biological molecules by label-free imaging techniques such as CARS microscopy and SRS microscopy (Mansfield et al., 2013). The C–F bond itself can be exploited to visualize the distribution of PFC in biological material. In this investigation, we have exploited this distinction to generate contrast images of the "negative space" within the mesophyll. Images of the apoplast throughout development will enable a better understanding of leaf expansion and growth. Moreover, is will be possible to use such techniques to develop refined models of the airspaces within leaves, and better understand the constraints of gaseous fluxes within.

## **CONCLUSION**

We have shown that mounting living leaves in PFD and perfluoroperhydrophenanthrene (PP11) improves image resolution for a number of laser-based microscopy techniques including LSCM, TPF, SHG, and SRS microscopy. These compounds allow greater *z-*axis penetration, resulting in clearer micrographs. In addition, PFCs may be used as label-free contrast agents to image the internal architecture of leaves, and enable a more precise understanding of the structural changes that occur during leaf development. As the mesophyll is a primary target for pathogenesis in plants, this technique may also be used to observe the processes of infection deep within the mesophyll. More generally, the use of PFCs as mounting media may be applied to other tissues rich in airspaces, such as animal lungs.

## **AUTHOR CONTRIBUTIONS**

George R. Littlejohn, Mark D. Fricker, Murray R. Grant, Nicholas Smirnoff, Julian Moger, Jacqueline T. Christmas, Richard M. Everson, and John Love designed the research; George R. Littlejohn, Eleanor Witterick, Jessica C. Mansfield, and Julian Moger performed the experiments; George R. Littlejohn, Eleanor Witterick, Jessica C. Mansfield, Jacqueline T. Christmas, Julian Moger, and John Love analyzed the data; George R. Littlejohn, Jessica C. Mansfield, Jacqueline T. Christmas and Jessica C. Mansfield and John Love wrote the manuscript.

## **ACKNOWLEDGMENTS**

The authors would like to thank John Mansfield and Dawn Arnold for providing GFP-labeled *Pseudomonas*. Funding for this research was from the Biotechnology and Biological Sciences Research Council (BBSRC), UK. We also acknowledge the Exeter Imaging Network.

#### **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 21 December 2013; accepted: 24 March 2014; published online: 23 April 2014.*

*Citation: Littlejohn GR, Mansfield JC, Christmas JT, Witterick E, Fricker MD, Grant MR, Smirnoff N, Everson RM, Moger J and Love J (2014) An update: improvements in imaging perfluorocarbon-mounted plant leaves with implications for studies of plant pathology, physiology, development and cell biology. Front. Plant Sci. 5:140. doi: 10.3389/fpls.2014.00140*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Littlejohn, Mansfield, Christmas,Witterick, Fricker, Grant, Smirnoff, Everson, Moger and Love. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Arabidopsis myosin XI sub-domains homologous to the yeast myo2p organelle inheritance sub-domain target subcellular structures in plant cells

#### *Amirali Sattarzadeh1,2, Elmon Schmelzer and Maureen R. Hanson1 2 \**

*1 Department of Molecular Biology and Genetics, Cornell University, Ithaca, NY, USA*

*2 Central Microscopy, Max-Planck-Institute for Plant Breeding Research, Cologne, Germany*

#### *Edited by:*

*Alex Costa, University of Milan, Italy*

*Reviewed by: Sherryl Bisgrove, Simon Fraser University, Canada Cristina Ruberti, Michigan State University, USA*

#### *\*Correspondence:*

*Elmon Schmelzer, Central Microscopy Max Planck Institute for Plant Breeding Research, Carl-von-Linne-Weg 10, 50829 Köln, Germany e-mail<EMAIL_ADDRESS>
Myosin XI motor proteins transport plant organelles on the actin cytoskeleton. The Arabidopsis gene family that encodes myosin XI has 13 members, 12 of which have sub-domains within the tail region that are homologous to well-characterized cargo-binding domains in the yeast myosin V myo2p. Little is presently known about the cargo-binding domains of plant myosin XIs. Prior experiments in which most or all of the tail regions of myosin XIs have been fused to yellow fluorescent protein (YFP) and transiently expressed have often not resulted in fluorescent labeling of plant organelles. We identified 42 amino-acid regions within 12 Arabidopsis myosin XIs that are homologous to the yeast myo2p tail region known to be essential for vacuole and mitochondrial inheritance. A YFP fusion of the yeast region expressed in plants did not label tonoplasts or mitochondria. We investigated whether the homologous Arabidopsis regions, termed by us the "PAL" sub-domain, could associate with subcellular structures following transient expression of fusions with YFP in *Nicotiana benthamiana*. Seven YFP::PAL sub-domain fusions decorated Golgi and six were localized to mitochondria. In general, the myosin XI PAL sub-domains labeled organelles whose motility had previously been observed to be affected by mutagenesis or dominant negative assays with the respective myosins. Simultaneous transient expression of the PAL sub-domains of myosin XI-H, XI-I, and XI-K resulted in inhibition of movement of mitochondria and Golgi.

**Keywords: yeast myo2p, myosin V, transient expression, Golgi, mitochondria, vacuole, confocal microscopy,** *Nicotiana benthamiana*

## **INTRODUCTION**

Myosins, molecular motors that move cargo along actin filaments, occur in all eukaryotic organisms. Based on sequence homology, the myosin super family can be divided into 37 classes (Richards and Cavalier-Smith, 2005; Foth et al., 2006). The myosins from higher plants fall within only two classes: class VIII and XI (Hodge and Cope, 2000; Berg et al., 2001). Plant myosins have the domain pattern that is characteristic for most of the myosins: (1) the extremely conserved N-terminal head (motor) domain, which binds to actin filaments and is responsible for force production via ATP hydrolysis (2) the neck (lever arm) domain, including distinctive repeat motifs called IQ repeats and (3) the C-terminal tail domain that facilitates cargo binding. Like most myosins, the tails of plant myosins are highly divergent domains that vary widely in length and in sequence. In addition, the tails of plant myosins have coiled-coil-forming sequences that allow the molecules to dimerize, producing two-headed molecules. In general, the tail domains of myosins are believed to be largely responsible for class-specific functions.

ATM1, which is a class VIII myosin, was the first plant myosin to be characterized, and was implicated in *Arabidopsis thaliana* cell wall formation (Reichelt et al., 1999). Recently, class VIII myosins were shown to be involved in plasmodesmata- mediated intracellular trafficking and endocytosis (Avisar et al., 2008a; Golomb et al., 2008; Sattarzadeh et al., 2008). In contrast to class VIII myosins, class XI myosins are similar to the wellstudied myosin V proteins of fungi and animals. In Arabidopsis, a 13-member gene family encodes the myosin XIs (Reddy, 2001; Tominaga and Nakano, 2012). Analysis of insertional mutants of the Arabidopsis myosin XIs revealed considerable redundancy of function, as little effect on plant growth and organelle dynamics was detected (Ojangu et al., 2007; Peremyslov et al., 2008). Single and higher order mutants and RNA silencing have been used to probe functions of the myosin XI family (Ojangu et al., 2007; Prokhnevsky et al., 2008; Sparkes et al., 2008; Avisar et al., 2009).  Replacing the motor domain with fluorescent proteins has resulted in mixed success regarding labeling of particular organelles. Depending on the region of the myosin that has been expressed, sometimes minimal labeling of subcellular structures have been observed, with most fluorescence remaining in the cytoplasm (Reisen and Hanson, 2007; Avisar et al., 2008b, 2009, 2012). The particular portion of the myosin XI tail that has been used in such experiments has a strong effect on whether or not fluorescent vesicles and subcellular structures are observed.

Class V myosin tails such as yeast myo2p contain two functional domains which harbor vacuole- and mitochondrialspecific and secretory vesicle-specific regions (Pashkova et al., 2005, 2006; Fortsch et al., 2011). Regions homologous to the two domains exist in the Arabidopsis myosin XI family. Previously we have described the labeling of subcellular locations by YFP fusions to the DIL (dilute) domain (similar to the yeast secretory vesiclespecific domain) of 12 Arabidopsis myosin XIs (Sattarzadeh et al., 2011). Peroxisomes were labeled by YFP fusions to DIL domains of 8 different myosin XIs, three labeled Golgi, and others labeled the plasma membrane, endoplasmic reticulum (ER), nuclear envelope, and unidentified vesicles, but none interacted with mitochondria (Sattarzadeh et al., 2011). Inspection of the members of the Arabidopsis myosin XI gene family revealed that except for XI-J, Arabidopsis myosin XI genes encode a 42 amino-acid region within the tail region that is homologous to the yeast myo2p region known to affect vacuole and mitochondrial inheritance (**Figure 1**). All of these regions in Arabidopsis exhibit the sequence PAL in this region; therefore we will refer to these 42 amino acids as the Arabidopsis myosin XI PAL sub-domains. We have previously expressed a YFP fusion to the PAL sub-domain of myosin XI-F and have shown that it localizes to plastids, stromules, peroxisomes, and Golgi in particular cells (Sattarzadeh et al., 2009). Here we report that YFP fusions of the PAL domain from 11 additional Arabidopsis myosin XIs localize to Golgi, mitochondria, nuclear envelope, the plasma membrane and/or to unidentified vesicles. While Golgi and mitochondrial mobility continued in cells transiently expressing the PAL sub-domains of either myosin XI-H, XI-I, and XI-K, simultaneous expression of all three PAL sub-domains resulted in cells with minimal or negligible movement of the two organelles.

## **RESULTS**

## **ARABIDOPSIS MYOSIN XI TAIL DOMAIN HOMOLOGOUS TO A YEAST MYOSIN V DOMAIN**

In the yeast myosin ScMyo2p, there is a 11 amino acid section that has been shown by mutagenesis to be critical for vacuole inheritance (Catlett et al., 2000). Mutations in this region also affect mitochondrial inheritance (Fortsch et al., 2011). A 42 amino acid region encompassing the 11 amino acids is highly conserved between yeast and plants and among the 12 different plant myosin XIs (**Figure 1B**). Based on the ScMyo2p structure template (PDB code2F6HX), homology models of globular tails of all *A. thaliana* class XI myosins were built. The three- dimensional structures that were obtained indicate that the predicted general architecture of the PAL sub-domain in class XI myosins is very similar to that of ScMyo2p (**Figure S1**; **Figure 1C**). The models indicate that the 11 amino-acid sections homologous to the corresponding yeast sections are predicted to be exposed on the surface of the polypeptides when the 42-amino acid regions we expressed are modeled.

#### **LOCALIZATION OF YFP::AT MYOSIN XI PAL SUB-DOMAIN FUSIONS**

Previously we have fused the PAL sub-domain from Arabidopsis myosin XI-F to YFP and have shown that the fusion protein labels plastids and stromules (Sattarzadeh et al., 2009). We have now made constructs with the remaining 11 PAL sub-domains from the Arabidopsis class XI myosins fused to YFP at the N-terminus. We report YFP fusion information for XI-C and XI-E together because the PAL domains of these two myosin XIs are identical. The localization of the YFP fusion proteins was examined by confocal laser scanning microscopy (CLSM) in leaf epidermal cells of *N. benthamiana* following agroinfiltration.

Except for the YFP::At XI-2 PAL fusion, all tested fusions labeled vesicles in the same size range as the Golgi, endosomes, and mitochondria (**Figures 2A–E**). The YFP fusions with myosin XI-2 failed to localize to any vesicular structures and strongly labeled the plasma membrane (**Figure 2A**; **Figure S2**). The YFP fusion with the At XI-A PAL sub-domain was usually seen also in the cytoplasm; however, some unidentified vesicular structures were sometimes observed in a few cells (**Figure 2B**). YFP fusion with the homologous region in yeast (RVI: Required for Vacuole Inheritance) resulted in diffuse cytoplasmic labeling in plant cells, but some punctate loci could be discerned at the plasma membrane (**Figure 2F**).

In order to identify the YFP-labeled organelles, we performed co-localization studies by co-expressing the myosin YFP-PAL fusions transiently with CFP or mCherry organelle markers in leaf epidermal cells of *N. benthamiana*. The YFP fusions with the PAL sub-domain of myosins XI-1, XI-D, XI-G, XI-H, XI-I and XI-K co-localized with the Golgi marker (**Figure 3**). Some but not all Golgi were labeled by the YFP::PAL fusion of myosin XI-B (**Figures 3D–F**). Vesicles labeled by YFP fusions with the PAL sub-domain of XI-C and XI-E did not coincide with the Golgi marker (data not shown).

YFP fusions with the PAL sub-domain from myosins XI-C, XI-E, XI-G, XI-H, XI-I, and XI-K co-localized with a mitochondrial marker (**Figure 4**). YFP::PAL fusions with myosins XI-B (**Figure 5**) and XI-D (data not shown) did not coincide with the mitochondrial signal. When YFP::PAL fusions of myosin XI-B, XI-H, XI-I, and XI-K were expressed along with a peroxisomal marker, no co-localization was observed (**Figure 5**).

Eight of the YFP fusions to PAL sub-domain (AtXI-1,B,C,D,E,G,I and K) appear to label the nuclear envelope in epidermal cells (**Figures 6A–I**). Even though the homologous region in yeast interacts with vacuoles, we observed only one plant PAL sub-domain, from XI-B, that label structures that are characteristic of tonoplast membranes in appearance (Xu et al., 2006) (**Figure 7**). Most cells expressing XI-B exhibited fluorescent Golgi, vesicles, and nuclear envelopes (**Figure 7**). The positive and negative co-localizations are summarized in **Table 1**.

## **MOTILITY OF THE VESICLES LABELED WITH YFP::PAL SUB-DOMAIN FUSIONS**

High levels of expression are often achieved following agroinfiltration. When a myosin tail domain lacking the motor function is expressed, a dominant negative effect on motility can sometimes be detected. However, when we examined the vesicles labeled by the 11 YFP::At PAL sub-domains, we found that rapid motility

was retained, in contrast to our prior observation of stationary vesicles following agroinfiltration of YFP::At DIL domain fusions. Motility is documented in **Figures 8A–C** and **Movies S1**– **S4** in Supplementary Material. In order to determine whether a dominant negative effect on movement might occur if multiple PAL sub-domains were expressed, we agroinfiltrated simultaneously four strains, containing YFP-PAL sub-domain fusions from myosins XI-H, XI-I, and XI-K, along with a strain carrying either a mCherry marker for Golgi or for mitochondria. About a third of the cells expressing YFP exhibited YFP-labeled vesicles that were stationery or poorly motile (**Figures 8D–F**). Agroinfiltration does not result in all cells in a leaf expressing the transgenes, so some cells may be expressing only one or two of the PAL subdomain fusions. We show an example of two adjacent cells, one of which is expressing one or more YFP-labeled PAL sub-domains localized to nearly immobile vesicles, while an adjacent cell is

expressing only the mCherry mitochondrial marker and therefore exhibits motile mitochondria (**Movie S5**). We provide an example of a cell expressing mCherry-Golgi and YFP:: PAL sub-domains located on immobile vesicles in **Movie S6**. When the mCherry Golgi marker is infiltrated by itself, the Golgi exhibit high motility (**Movie S7**). We also observed minimal motility of YFP-labeled vesicles in many cells when we co-infiltrated three strains containing YFP-PAL sub-domain fusions from myosins XI-H, XI-I, and XI-K, without simultaneous co-infiltration of a strain carrying an organelle marker (**Movie S8**). In **Figure 8**, we provide three panels (A–C) from a movie that illustrate movement of vesicles labeled with YFP:: XI-K PAL when it is expressed alone; however, in (**Figures 8D–F)**, no movement of vesicles occurs following coinfiltration of XI-H, XI-I, and XI-K. We conclude that expression of multiple PAL sub-domains can impair movement of Golgi and mitochondria by a dominant negative effect.

## **DISCUSSION**

We have observed that 42-amino acid portions of 12 Arabidopsis myosin XI tail regions (the PAL sub-domain) are targeted to different subcellular organelles and membranes. Previously Li and Nebenfuhr (2007) expressed larger regions of the Arabidopsis XI-1 and XI-2 tails that encompass the PAL sub-domain and found they were targeted to peroxisomes, unidentified vesicles, or remained in the cytoplasm (**Figure 9**). We recently reported that the DIL domain of Arabidopsis myosin XIs, which is Cterminal to the PAL sub-domain and also encompassed by the constructs used by Li and Nebenfuhr (2007), targets YFP to several organelles and membranes (Sattarzadeh et al., 2011). PAL and DIL regions from particular myosin XIs can target YFP to Golgi, nuclear envelope, and/or plasma membrane. However, no DIL domains resulted in localization of YFP to mitochondria or tonoplast. Conversely, none of the PAL sub-domain fusions resulted in labeling of ER nor peroxisomes, while 8 DIL domains targeted YFP to peroxisomes and two labeled ER (Sattarzadeh et al., 2011). Thus there are some general differences in the localization information provided by the two different tail regions.

Although the PAL sub-domains among the *A. thaliana* class XI myosins are quite similar, there are some sequence differences in this 42 amino acid region of individual tail domains that could explain the different subcellular localizations (**Table 2**). While the PAL sub- domains of XI-C and XI-E are completely identical, other myosin XI PAL regions are less similar; nevertheless, it is not possible to consult the table of identity to predict which PAL subdomains are likely to label one organelle vs. another. For example, of the PAL sub-domains that label mitochondria, XI-C/E PAL is 71, 80, 85, and 95% identical, respectively, to XI-G, XI-H, XI-I, and XI-K PAL. However, XI-C/E PAL is 85, 83, and 85%, identical, respectively to XI-1, XI-B, and XI-F, which do not label mitochondria but do label Golgi (**Table 2**). Unidentified aspects of the protein tertiary structure may be important for selective labeling.

The PAL sub-domain of the myosin XIs was selected for study because it is homologous to a sub-domain in yeast myo2p required for vacuole inheritance (Catlett et al., 2000). Recently, (Fortsch et al., 2011) have shown that the same yeast myo2p region is important for mitochondrial inheritance. Furthermore, Li and Nebenfuhr (2007) demonstrated that the structure of myosin XI-1 could be modeled with structural data available for yeast myo2p. Ancestral myosin XIs may have duplicated and diverged in the tail domains to interact with a variety of subcellular structures, including mitochondria, Golgi, and peroxisomes. The yeast PAL sub-domain did not label plant mitochondria or tonoplast (**Figure 2F**), though the yeast DIL domain was able to label plant peroxisomes (Sattarzadeh et al., 2011).

High-level expression of defective motor proteins in various organisms have often been observed to create a dominantnegative effect on organelle movement, possibly because saturation of receptors with the motorless protein prevents intact motors from binding. When we transiently expressed YFP::DIL fusions of 12 Arabidopsis myosin XI proteins, we found that

**and mCherry markers for Golgi in epidermal leaf cells of** *N. benthamiana* **(A) At XI-1. (D)** At XI-I B. **(G)** At XI-D. **(J)** At XI-G. **(M)** At XI-H. **(P)** At XI-I **(S)** At XI-K. **(B,E,H,K,N,Q,T)** Golgi marker. **(C,F,I,L,O,R,U**) Merged images of YFP (yellow) and mCherry (red) or CFP (blue). Scale bar = 50μm

expression of all but one of the fusions resulted in loss of motility of YFP-labeled vesicles and organelles (Sattarzadeh et al., 2011). Expression of various portions of fluorescent protein fusions to myosin XI tails have sometimes resulted in detectable effects on motility even though the affected organelles were not visibly fluorescently labeled. For example, Avisar et al. (2009) did not document localization of GFP::myosin XI tail regions to Golgi or mitochondria, but did detect inhibition of movement of both

**with mCherry markers for mitochondria in epidermal leaf cells of** *N. benthamiana***.  Scale bar = 50μm.

of the organelles when fusions to XI-1, XI-2, XI-C, XI-E, XI-I, and XI-K were expressed in *N. benthamiana.* Vick and Nebenfuhr (2012) have pointed out that expression of myosin tail regions might result in non-specific inhibition of motility as well as receptor-specific inhibition. We detected no Golgi or mitochondrial labeling with either the PAL or DIL regions of XI-2, but YFP::PAL of XI-C and E labeled mitochondria, YFP: DIL of XI-E labeled peroxisomes and YFP::DIL of XI-1 labeled Golgi (**Figure 3** and Sattarzadeh et al., 2011).

We have expressed Arabidopsis PAL domains in a heterologous plant system, namely *N. benthamiana*. The PAL domains in this species are highly similar to those in Arabidopsis (**Figure 1**; **Figure S3**). In fact, there are *N. benthamiana* PAL domains that are identical to those in Arabidopsis XI-K and XI-C/E (**Figure S3**). Thus proteins present in *N. benthamiana* that bind to PAL domains of endogenous myosin XIs are likely to be able to interact strongly with expressed Arabidopsis PAL domains. However, divergence between some of the Arabidopsis and Nicotiana PAL domains may result in weaker or absent interaction with Nicotiana endogenous proteins, potentially explaining why expression of a single Arabidopsis PAL domain is not sufficient for a strong dominantnegative effect. Furthermore, we may not have detected all the subcellular structures that interact with Arabidopsis PAL domains in Arabidopsis, if they are so divergent that they cannot interact with the myosin receptor proteins in Nicotiana.

Sparkes et al. (2008) were able to observe fluorescent puncta when YFP fusions to tail regions of Arabidopsis XI-E and XI-K were expressed in *N. tabacum* epidermal cells. Although not all peroxisomes, mitochondria, and Golgi were labeled by the YFP fusions to XI-E and XI-K, inhibition of their movement was detected in the cells (Sparkes et al., 2008). We observed that the YFP::PAL sub-domain fusions of XI-I and XI-K labeled mitochondria and Golgi in *N. benthamiana*, but YFP::PAL XI-H also labeled both organelles. Our YFP::PAL XI-B, XI-F, and XI-H fusions labeled Golgi. This is consistent with the observations of Avisar et al. (2009), who observed inhibition of Golgi movement by fusions with XI-B, XI-F, and XI-H; however, the inhibition occurred only in *N. tabacum* and not in

*N. benthamiana*. Thus if a PAL sub-domain labels a particular motile organelle, the larger constructs expressed by others have often resulted in effects on motility of that organelle. But the correlation is not perfect, as we did not detect labeling of all organelles whose movement was affected by expression of a particular myosin tail. For example, although Sparkes et al. (2008) observed inhibition of Golgi by expression of YFP::XI-E tail fusions, neither the PAL nor DIL regions of XI-E labeled Golgi, even though the XI-E PAL sub-domains in *A. thaliana* and *N. benthamiana* are identical. Possibly other sequences we have not analyzed are able to interact with Golgi and mitochondria; however, abnormal conformation of myosin fragments may also prevent interactions that would normally occur with the intact myosin.

Single myosin XI insertional mutants do not exhibit major phenotypes, though XI-2 and XI-K knockouts are affected in root hair elongation and in Golgi, mitochondrial, and peroxisome motility (Peremyslov et al., 2008). While we have not been able to observe labeling of these three organelles with YFP fusions with XI-2 PAL sub-domains, all three organelles are labeled by either the PAL sub-domain (mitochondria, Golgi) or the DIL domain (peroxisomes) of XI-K fused to YFP. In plants containing single knockouts of XI-1, XI-2, XI-C, XI-E, and XI-I, Avisar et al. (2012) detected minor effects on Golgi movement, while an XI-K knockout greatly inhibited Golgi motility. Of these myosins, we found that YFP::PAL sub-domain fusions of XI-1, XI-I, and XI-K and the YFP::DIL fusion of XI-C labeled Golgi (Sattarzadeh et al., 2011).

Of the Arabidopsis myosin XIs, XI-1, XI-2, XI-H, XI-I, and XI-K are the most highly and ubiquitously expressed (Peremyslov et al., 2011; Sparkes, 2011). Much attention has been focused on myosin XI-K. Avisar et al. (2008b) observed that transient

**myosin PAL sub-domains in epidermal leaf cells of** *N. benthamiana***.** YFP fusions of At XI-1 PAL **(A)**, At XI-B PAL **(B)**, At XI-C/E PAL **(C)**, At XI-D PAL **(D)**, At staining of epidermal leaf cells of N. benthamiana expressing YFP:: At XI-K PAL. **(I)** Merged images of YFP (yellow) and DAPI staining (blue). Scale bar = 15μm.

silencing or expression of dominant negative constructs of myosin XI-K resulted in impaired movement of Golgi, peroxisomes, and mitochondria. More recent experiments with the XI-K tail region of *N. benthamiana* demonstrated that the particular myosin fragment that is expressed greatly influences how much vesicle motility is affected (Avisar et al., 2012). When Avisar et al. (2012) expressed two globular tail regions that encompass the corresponding Arabidopsis XI-K PAL and DIL domains that we have utilized, they observed inhibition of Golgi movement, but a third construct with a C-terminal extension, and two single aminoacid mutations in the DIL domain resulted in retention of Golgi motility (Avisar et al., 2012). When the entire XI-K sequence was fused to YFP, a small fraction of the Golgi, some regions of the ER, and many endomembrane vesicles were labeled (Peremyslov et al., 2012). The importance of myosin XI-K is also indicated in our YFP fusion experiments, as XI-K PAL or DIL regions fused to YFP resulted in labeling of many locations: Golgi, mitochondria, peroxisomes, nuclear envelope, plasma membrane, and ER (**Figures 3**, **4**; Sattarzadeh et al., 2011).

Even though expression of 12 different Arabidopsis YFP::PAL sub-domain fusions resulted in readily detectable fluorescent labeling of various subcellular structures, the labeled Golgi and mitochondria move rapidly (**Movies S1**–**S4**). We have identified 6 different myosin XIs whose PAL sub-domains can interact with mitochondria, and 7 myosin XIs whose PAL sub-domains can interact with Golgi. When certain tail regions of 6 myosin XIs

**Table 1 | Summary of localization of PAL sub-domains of** *A. thaliana* **class XI myosins N-terminally fused to YFP and transiently expressed in leaves epidermal cells of** *N. benthamiana***.**

*Organelles that were observed not to co-localize with a particular myosin are shown in italics.*

were labeled in transient assays, we previously observed that the motility of vesicles was retained (Reisen and Hanson, 2007). In contrast, when we expressed YFP::DIL fusions in *N. benthamiana*, movement of Golgi and peroxisomes ceased (Sattarzadeh et al., 2011). We cannot rule out the possibility that some of the targeting of YFP::PAL sub-domains to organelles may be due to chance function of the 42 amino-acid sequence as a transit peptide or secretory signal, as C-terminal fusions can sometimes unexpectedly result in labeling of peroxisomes and other vesicles (Cutler et al., 2000). However, targeting programs do not predict any of the PAL sub-domains to be effective transit or signal sequences (**Table S1** in Supplementary Material), though we recognize that such predictions are not perfect. The organelles labeled by the myosin XI PAL sub-domains are largely consistent with a role for the individual myosin XIs in organelle motility, according to the results of mutant, silencing, and dominant negative analysis. Possibly the YFP::PAL sub- domains do not accumulate to the same levels as the YFP::DIL fusion, thus not saturating receptors, allowing a sufficient number of intact motors to attach and transport the organelles. Alternatively, there may be differences in the Golgi and mitochondrial receptors that interact with the multiple PAL sub-domains, so that even if one receptor is saturated with a defective myosin, other receptors can interact with other myosins. Different myosin XIs may vary in affinity for particular cargoes, especially in a heterologous system. Less than 7% of the Golgi were labeled with a full-length myosin XI-K::YFP fusion (Peremyslov et al., 2012). We did not observe a dominant negative effect on Golgi and mitochondrial motility until we expressed three PAL sub-domains (XI-H, XI-I, XI-K) simultaneously (**Movies S5**, **S6**, **S8**). Further inquiry into the role of the PAL sub-domains is needed in order to determine whether motility of Golgi and mitochondria is affected as a result of saturation of receptors on the cargo or because of unknown non-specific effects on myosin function.

**FIGURE 8 | Sequential images indicating movement YFP::At XI-K PAL sub-domain-labeled organelles but lack of movement when multiple domains are infiltrated simultaneously. (A–C)** *N. benthamiana* leaves were infiltrated with an Agrobacterium strain carrying a YFP::At XI-K PAL sub-domain. **(D–F)** Leaves were infiltrated simultaneously with three

Agrobacterium strains (YFP::At XI-H, YFP::At XI-I, and YFP::At XI-K PAL sub-domains), each of which contained a transgene with one PAL sub-domain fused to YFP. Time-lapse series were obtained by confocal microscope. Only three frames are shown. Time (*t*) is indicated in seconds. Bar = 50 um. An arrow marks an organelle that is in different positions in **A–C**.

**Table 2 | Pairwise comparison of the amino acid sequences of the PAL sub-domain from** *A. thaliana* **class XI myosins and vacuole binding site in the yeast class V myosin Myo2p.**

*The identity is shown in percent and the number of identical amino acids are listed.*

**FIGURE 9 | Diagram of At XI-K constructs that have been expressed in Nicotiana.** Constructs diagramed are (1) from Sattarzadeh et al. (2011), (2) from Reisen and Hanson (2007), (3) from Li and Nebenfuhr (2007), (4) from Sparkes et al. (2008), and (5) from Avisar et al. (2009). CC: coiled-coil. IQ: IQ repeats. BiFC: construct for Bimolecular Fluorescence Complementation. PM: Plasma Membrane.

## **MATERIALS AND METHODS**

## **GENERATION OF YFP::MYOSIN CONSTRUCTS, PLANT GROWTH AND AGROINFILTRATION**

Generation of the YFP::myosin constructs was performed as described (Sattarzadeh et al., 2009) using standard TOPO and Gateway cloning system (*Invitrogen*, Carlsbad, CA). The primers used are listed in **Table S2** in Supplementary Material. Plant growth and agroinfiltration was done as described (Sattarzadeh et al., 2011). When multiple strains were infiltrated, the same total concentration of bacteria was used as for single infiltration.

## **FLUORESCENT LABELING OF ORGANELLES AND CONFOCAL LASER SCANNING MICROSCOPY**

For labeling of mitochondria, peroxisomes, Golgi stacks and plasma membrane with mCherry fluorescent protein, binary vectors, CD3-991, CD3-984, CD3-967, and CD3-1008 (Nelson et al., 2007) were used, respectively. According to Nelson et al. (2007), the mitochondrial marker was made by fusing the first 29 amino acids from the N-terminus of yeast COXIV (Köhler et al., 1997) to mCherry, the peroxisomal targeting signal Ser-Lys-Leu (Reumann, 2004) to mCherry, and the Golgi marker resulted from fusion of the first 49 amino acids of soybean α-1,2-mannosidase I (Saint-Jore-Dupas et al., 2006) to mCherry. The Golgi CFP marker was made by Sattarzadeh et al. (2011) by fusing the first 52 N-terminal amino acids of a rat 2, 6-sialyl transferase (Boevink et al., 1998). For DNA staining by 4- ,6 diamidino-2-phenylindole (DAPI), leaves were immersed in a solution of 2.5% mannitol/0.01%silwet/300 nM DAPI.

CSLM was performed on a Leica microscope equipped with a TCS-SP2 confocal scanning head (Leica Microsystems, Heidelberg, Germany) as described (Sattarzadeh et al., 2009). **Movies S5**–**S7** were captured by using an Andor XD revolution spinning disc confocal microscope. Infiltrations of each YFP::PAL fusion were performed at 3 separate times and approximately 100 cells in total were observed for each YFP::PAL fusion. The images shown represent the appearance of the majority of the labeling patterns for each fusion. We note in the text if an image is provided of labeling pattern that was only rarely observed.

## **BIOINFORMATIC ANALYSIS AND STRUCTURAL HOMOLOGY MODEL ANALYSIS**

GeneDoc was used for generation of alignments (Nicholas and Nicholas, 1997). Percent identity and divergence between Arabidopsis and *N. benthamiana* myosin XI PAL domains was generated with the LASERGENE MEGALIGN program (DNAstar, Madison, WI). Structure homology models for the tail domain of class XI mysoins were built by SWISSMODEL (Arnold et al., 2006) based on the Myo2p structure template (2f6hX). PyMol (Delano Scientific) was used for drawing the structural homology models.

## **ACKNOWLEDGMENTS**

This work was supported by the Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, U.S. Department of Energy grant DE-FG02-09ER16070 to Maureen R. Hanson. We thank Rainer Franzen for excellent technical assistance and Rebecca Williams at the Cornell Life Sciences Core Laboratories Center for Microscopy and Imaging for numerous helpful suggestions regarding the spinning disc confocal microscope.

## **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www.frontiersin.org/Journal/10.3389/fpls.2013.00407/ abstract

**Supplementary Figure 1 | Homology structural modeling of myosin XI globular tail based on similarity to the myosin V ScMyo2p.** Predicted three-dimensional structures of the globular tail of *A. thaliana* class XI myosins are illustrated. **(A)** XI-1. **(B**) XI-2. **(C)** XI-A. **(D)** XI-B. **(E)** XI-C. (**F**) XI-D. **(G)** XI-E. **(H)** XI-F. **(I)** XI-G. **(J)** XI-H. **(K)** XI-I. **(L)** XI-K. Surface residues are shown in orange. The 11 amino acids critical for the vacuole binding site in ScMyo2p tail (Catlett et al., 2000) and the corresponding amino acids in PAL sub-domains are shown in cyan.

**Supplementary Figure 2 | Co-expression of YFP::Myosin XI-2 PAL domain with mCherry markers for the plasma membrane in epidermal leaf cells of** *N. benthamiana* **(A) At XI-2 (B) Plasma membrane marker. (C)** Merged images of YFP (yellow) and mCherry (red). Scale bar = 10μm.

## **Supplementary Figure 3 | Pairwise comparison of the amino acid sequences identity and divergence between Arabidopsis and**

*N. benthamiana* **myosin XI PAL domains.** Table of identity and divergence was generated with the LASERGENE MEGALIGN program(DNAstar, Madison, WI).

#### **Movies S1–S4 | Movement of YFP::At XI-PAL sub-domain-labeled**

**organelles.** *N. benthamiana* leaves were infiltrated with an Agrobacterium strain carrying a YFP::AtXI-PAL sub-domain **(S1)** XI-1 **(S2)** XI-C/E **(S3)** XI-D and **(S4)** XI-K.

**Movie S5 | Co-expression of YFP::At XI-H, YFP::At XI-I and YFP::AT XI-K PAL sub-domains with mCherry markers) for mitochondria in epidermal leaf cells of** *N. benthamiana* **leaves.** Leaves were infiltrated simultaneously with four Agrobacterium strains, each of which contained either contained

## **REFERENCES**

a transgene with one PAL sub-domain fused to YFP (yellow) or a mCherry protein (red) targeted to mitochondria. Time-lapse series (25 frames, 0.2 s intervals) were obtained with an Andor spinning disc confocal microscope using a 60X objective. To the right of the yellow-labeled cell is a cell with motile mCherry-labeled mitochondria that is not exhibiting expression of the YFP-labeled PAL sub-domains.

**Movie S6 | Co-expression of YFP::At XI-H, YFP::At XI-I and YFP::AT XI-K PAL sub-domains with mCherry markers (red) for Golgi in epidermal leaf cells of** *N. benthamiana* **leaves.** Leaves were infiltrated simultaneously with four Agrobacterium strains, each of which contained either contained a transgene with one PAL sub-domain fused to YFP (yellow) or a mCherry protein (red) targeted to mitochondria. Time-lapse series (25 frames, 0.2 s intervals) were obtained with an Andor spinning disc confocal microscope using a 60X objective.

**Movie S7 | Movement of mCherry-labeled Golgi bodies in epidermal cells of** *N. benthamiana* **leaves.** Time-lapse series (20 frames, 3 s intervals) were obtained by the Leica CSLM. The movie is shown at 5 frames per second.

**Movie S8 | Co-expression of YFP::At XI-H, YFP::At XI-I and YFP::AT XI-K PAL sub-domains in epidermal leaf cells of** *N. benthamiana* **leaves.** Leaves were infiltrated simultaneously with three Agrobacterium strains, each of which contained a transgene with one PAL sub-domain fused to YFP. Time-lapse series (25 frames, 0.2 s intervals) were obtained with an Andor spinning disc confocal microscope using a 60X objective.

**Table S1 | Predicted subcellular localization of** *A. thaliana* **class XI myosin PAL sub-domains.** The prediction was generated using Target P (Brunak et al., 2007) based on the predicted presence of any of the N-terminal presequences: chloroplast transit peptide (cTP), mitochondrial targeting peptide (mTP) or secretory pathway signal peptide (SP).

#### **Table S2 | List of primers used in this study.**

on an actin/ER network. *Plant J.* 15, 441–447. doi: 10.1046/j.1365- 313X.1998.00208.x

F-actin. *Front. Plant Sci.* 3:184. doi: 10.3389/fpls.2012.00184

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 01 July 2013; accepted: 26 September 2013; published online: 22 October 2013.*

*Citation: Sattarzadeh A, Schmelzer E and Hanson MR (2013) Arabidopsis myosin XI sub-domains homologous to the yeast myo2p organelle inheritance sub-domain target subcellular structures in plant cells. Front. Plant Sci. 4:407. doi: 10.3389/fpls.2013.00407*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2013 Sattarzadeh, Schmelzer and Hanson. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

**REVIEW ARTICLE** published: 19 November 2013 doi: 10.3389/fpls.2013.00439

## *Alexander Ivakov\* and Staffan Persson\**

*Max-Planck-Institute of Molecular Plant Physiology, Potsdam, Germany*

#### *Edited by:*

*Alex Costa, University of Milan, Italy*

#### *Reviewed by:*

*Laurie Smith, University of California San Diego, USA Ying Fu, China Agricultural University, China*

*\*Correspondence: Alexander Ivakov and Staffan Persson, Max-Planck-Institute of Molecular Plant Physiology, Am Muehlenberg 1, 14476 Potsdam, Germany e-mail: Ivakov@mpimp-golm. mpg.de;Persson@ mpimp-golm.mpg.de*

Plant cell shape, seen as an integrative output, is of considerable interest in various fields, such as cell wall research, cytoskeleton dynamics and biomechanics. In this review we summarize the current state of knowledge on cell shape formation in plants focusing on shape of simple cylindrical cells, as well as in complex multipolar cells such as leaf pavement cells and trichomes. We summarize established concepts as well as recent additions to the understanding of how cells construct cell walls of a given shape and the underlying processes. These processes include cell wall synthesis, activity of the actin and microtubule cytoskeletons, in particular their regulation by microtubule associated proteins, actin-related proteins, GTP'ases and their effectors, as well as the recently-elucidated roles of plant hormone signaling and vesicular membrane trafficking. We discuss some of the challenges in cell shape research with a particular emphasis on quantitative imaging and statistical analysis of shape in 2D and 3D, as well as novel developments in this area. Finally, we review recent examples of the use of novel imaging techniques and how they have contributed to our understanding of cell shape formation.

**Keywords: Microtubules, actin, auxin, pavement cell, shape tools, Arabidopsis, hypocotyl**

## **INTRODUCTION**

In immature organs where cell division is actively taking place, cells are small and the shapes of cells are largely defined by the most recent cell divisions where new cell walls have been laid down along previous division planes. The walls in such cells are straight and the cells are largely iso-diametric, bearing little resemblance to the shapes commonly observed in mature organs. To acquire their final shapes, cells must change their proportions while increasing in size. For changes in proportions to occur some parts of the cell surface must grow differently to other parts. This model of cell shape formation was proposed by Paul Green in 1965 on the basis of observations of the growth of giant internodal cells of *Nitella* and is still relevant today (Green, 1965).

In this review we aim to dissect how simple and complex cell shapes, such as those of cylindrical hypocotyl cells and leaf epidermal pavement cells, respectively, are thought to be established. A largely neglected area in the field of plant cell shape is the usage of quantitative means to define shape. We therefore also put an emphasis on how certain tools and algorithms may be used to quantify and compare shapes.

## **GENERAL CONCEPTS**

The shape of the cell is bounded and defined by the cell wall and it follows that cell wall expansion must be differentially regulated around the cell to generate the final cell morphology. Cell walls expand through irreversible deformation under a turgor-driven cell wall stress in a process called "creep." Creep involves the breaking of hydrogen bonds between cell wall polymers and the reversible and irreversible breakage of covalent bonds (Cosgrove, 2005). Cell wall expansion is promoted by cell wall loosening factors, such as expansin proteins, and xyloglucan endotransglycosylases/hydrolases (XTHs) (Cosgrove, 2000; Van Sandt et al., 2007). Expansins are thought to promote cell wall creep by breaking hydrogen bonds between cellulose and xyloglucan chains (Cosgrove, 2000). XTH is able to hydrolyse covalently-bonded xyloglucan chains and re-attach them at a later time, weakening the wall (Fry et al., 1992). Cell wall stiffening may also occur. Agents proposed to stimulate cell wall stiffening include pectin methylesterase (Micheli, 2001), extensin proteins, peroxidases (Passardi et al., 2004), and reactive oxygen species (Schopfer, 1996). Creep is typically measured in isolated cell walls in an extensiometer, where the wall is loaded with a constant load and the irreversible lengthening of the wall is measured (Cosgrove, 2000).

Differential cell wall expansion underlying shape formation can include growth rates varying locally as some parts of the wall expand faster than others, as well as growth anisotropy, where the rates of expansion of a given segment of the wall are different in different directions. Anistropy is distinguished from isotropy, where the rates of expansion are equal in all directions. Cell wall anisotropy is determined by the structure of the cell wall and the arrangement of cellulose within it. Cellulose consists of high molecular weight glucan chains arranged in partially-crystalline bundles held together by numerous hydrogen bonds. These structures, termed microfibrils, have a high tensile strength and strongly resist deformation in the presence of a stretching force. The strong microfibrils are embedded in a pliant gel-like matrix consisting of hemicelluloses and pectin polysaccharides. This fiber and matrix configuration of the cell wall has been likened to a composite material (e.g., fiber-glass) and cell walls have been shown to behave mechanically like such material (Kerstens et al., 2001; Fratzl et al., 2004). The orientation of the cellulose microfibrils within the cell wall is therefore an important determinant of the behavior of the wall during growth as well as in mature tissues (Burgert and Fratzl, 2009). Microfibrils resist expansion most strongly parallel to themselves and less strongly in other directions and thus form the basis for anisotropy.

Cellulose is produced by cellulose synthase (CesA) complexes which are mobile in the plasma membrane and extrude glucan chains into the cell wall. Cellulose microfibrils have long been observed to co-align with microtubules in the cell cortex (Green, 1962). More recently, observations of fluorescently-labeled CesA complexes in the plasma membrane showed that their movement is guided by cortical microtubules (Paredez et al., 2006). Hence, by controlling microtubule organization plant cells can control the arrangements of cellulose microfibrils, and therefore the pattern of wall expansion and cell morphogenesis (Wymer and Lloyd, 1996; Lloyd, 2011). Plant microtubules are mobile and highly dynamic structures which exhibit a treadmilling movement generated by a fast rate of net polymerization at the plus end and a slower rate of depolymerisation at the minus end (Shaw et al., 2003). Microtubules interacting with each other have emergent self-organizing properties and spontaneously organize into parallel bundles or arrays (Wasteneys and Ambrose, 2009). These microtubule arrays have the ability to rapidly re-organize to new orientations. Such re-organization control CesA trajectories and microfibril arrangements, which in turn influence cell shape.

## **CYLINDRICAL CELLS**

Anisotropic expansion has been shown to be responsible for the formation of shape in cylindrical cells, such as those of *Nitella* and *Chara*, but also of expanding root and hypocotyls cells in Arabidopsis. When cells in a multicellular organ grow in this manner, cylindrical organs such as stems, roots and hypocotyls are produced. Quantification of local strain rates in growing roots indicate a high degree of anisotropy in the expansion zone which contributes to the elongated cylindrical shape of the cells and of the entire organ and observations of microfibril orientations can largely explain the observed anisotropy (Baskin et al., 1999; Anderson et al., 2010).

## **CELLULOSE DEPOSITION**

Genetic and pharmacological experiments have shown that anisotropic expansion is greatly compromised in plants with decreased levels of cellulose (See Table S1 for an overview of mutants affecting cell shape). Cellulose-deficient mutants and plants treated with the cellulose synthesis inhibitor isoxaben exhibit radial swelling and dramatic decreases in cell elongation in hypocotyls and roots (Arioli et al., 1998; Fagard et al., 2000; Refrégier et al., 2004; Fujita et al., 2013). Presumably, in the presence of insufficient amounts of cellulose the cell wall is unable to resist the force of turgor and expands uncontrollably.

However, radial swelling is also sometimes observed in mutants with normal amounts of cellulose and with unaltered cellulose microfibril orientations, such as the temperature-sensitive microtubule organization mutant *mor1-1* (Sugimoto et al., 2003). Observations of this mutant led Wasteneys (2004) to propose the "microfibril length regulation hypothesis" (Wasteneys, 2004) whereby microtubules also control the length of deposited cellulose microfibrils. According to this hypothesis, in certain states of microtubule disruption, cellulose microfibrils are short and unable to resist the expansion of the cell wall due to insufficient hydrogen bonding with the cell wall matrix. These defects would then lead to radial swelling of the cells and decreased anisotropy.

## **MICROTUBULE ORGANIZATION**