Sphingolipids involvement in plant endomembrane differentiation: the BY2 case

Sphingolipids involvement in plant endomembranedifferentiation: the BY2 case

Anne Aubert1,†, Jessica Marion1,2,†, Claire Boulogne1, Mickael Bourge1,2, Susana Abreu1, Yannick Bellec3, Jean-Denis Faure3

and Beatrice Satiat-Jeunemaitre1,2,*

1Laboratoire Dynamique de la Compartimentation Cellulaire, CNRS UPR2355/IFR87, Institut des Sciences du Vegetal,

Centre de Recherche de Gif (FRC3115), 91198, Gif-sur-Yvette Cedex, France,2Pole de Biologie Cellulaire, Imagif, Centre de Recherche de Gif, (FRC3115), CNRS, IFR 87, 91198, Gif-sur-Yvette Cedex,

France, and3Institut Jean-Pierre Bourgin, unite mixte de recherche 1318, INRA-AgroParisTech, Centre de Versailles-Grignon,

78026 Versailles cedex, France

Received 14 October 2010; revised 10 December 2010; accepted 28 December 2010; published online 18 February 2011.*For correspondence (fax +33 (0)1 69 82 3355; e-mail [email protected]).†These authors have equally contributed to this work.

SUMMARY

Sphingolipids play an essential role in the functioning of the secretory pathway in eukaryotic organisms. Their

importance in the functional organization of plant cells has not been studied in any detail before. The

sphingolipid synthesis inhibitor fumonisin B1 (FB1), a mycotoxin acting as a specific inhibitor of ceramide

synthase, was tested for its effects on cell growth, cell polarity, cell shape, cell cycle and on the ultrastructure of

BY2 cells. We used cell lines expressing different GFP-tagged markers for plant cell compartments, as well as a

Golgi marker fused to the photoconvertible protein Kaede. Light and electron microscopy, combined with flow

cytometry, were applied to analyse the morphodynamics and architecture of compartments of the secretory

pathway. The results indicate that FB1 treatment had severe effects on cell growth and cell shape, and induced

a delay in cell division processes. The cell changes were accompanied by the formation of the endoplasmic

reticulum (ER)-derived tubular aggregates (FB1-induced compartments), together with an inhibition of cargo

transport from the ER to the Golgi apparatus. A change in polar localization of the auxin transporter PIN1 was

also observed, but endocytic processes were little affected. Electron microscopy studies confirmed that

molecular FB1 targets were distinct from brefeldin A (BFA) targets. We propose that the reported effects

of inhibition of ceramide biosynthesis reflect the importance of sphingolipids during cell growth and

establishment of cell polarity in higher plant cells, notably through their contribution to the functional

organization of the ER or its differentiation into distinct compartments.

Keywords: endomembrane, plant cells, light and electron microscopy, sphingolipids, fumonisin B1.

INTRODUCTION

The functional organization of eukaryotic cells by a distinct

set of endomembranous compartments is a key process in

evolution. The important roles of organelles such as the

endoplasmic reticulum (ER), Golgi apparatus (GA), endo-

somes and vacuoles/lysosomes in cell growth, cell division

and cell polarity processes include vectorial transport

activities along a cell axis from ER to plasma membrane, or

from plasma membrane into the cell, and imply specific

protein machineries (see references in Kepes et al., 2005).

Membrane lipids have been described to act both as struc-

tural components of the endomembranes and as signaling

molecules in the secretory pathway (Wolf et al., 1999;

Wisniewska et al., 2003; Raffaele et al., 2009b). Hence, it is

clear that they play a significant role in processes such as

subcellular compartmentation and transport/trafficking.

Various studies are currently under way to uncover the

functional organization of lipids in parts of the endomem-

brane systems in eukaryotic organisms. The general out-

come is a complex picture in which the biosynthetic steps of

membrane lipids involved different cell compartments.

Results also indicate that membrane lipids do not mix

homogenously, and that their interactions with other lipids

or proteins produce distinct domains within the membranes.

This spatial arrangement of membrane lipids in distinct

958 ª 2011 The AuthorsThe Plant Journal ª 2011 Blackwell Publishing Ltd

The Plant Journal (2011) 65, 958–971 doi: 10.1111/j.1365-313X.2011.04481.x

micro- or even nanodomains, rich in sphingolipids and

sterols, the so-called lipid rafts, are expected to play a role in

protein sorting throughout the endomembrane system in all

eukaryotic cells (Laloi et al., 2007; Klemm et al., 2009).

These main features of lipid compartmentation are also

shared by plant cells. However, plant cells also face specific

challenges with respect to environmental and evolutionary

conditions/constraints and, therefore, their endomembrane

system and lipid homeostasis exhibit specialized features

(respectively Hawes and Satiat-Jeunemaitre, 2005; Moreau

et al., 2007; Boutte et al., 2006; Brown et al., 2010). For

example, the lipid composition of the chloroplast mem-

brane, as well as lipid gradients from the ER to the plasma

membrane or vacuoles, was demonstrated to be plant-

specific using biochemical techniques (Yoshida and

Uemura, 1986; Moreau and Cassagne, 1994; Moreau et al.,

1998; Block et al., 2007). Hitherto the cell biology of plant

lipids, i.e. their in cellulo compartmentalized synthesis,

transport and subcellular interactions, have remained poorly

understood. The impact of phytosterols on the regulation

of cell polarity, cell division and on endomembrane organi-

zation has now been demonstrated in a number of elegant

studies (Willemsen et al., 2003; Boutte et al., 2007a). In

contrast, studies on the role of sphingolipids in cell

development are still in their infancy.

Compared with glycerolipids (i.e. phospholipids and

glycolipids), which form the main lipid constituents of the

plant endomembranes, sphingolipids and sterols are pres-

ent in relatively small quantities. The metabolism of sphin-

golipids in plants has many features in common with that

in other organisms. Ceramide is the basic backbone for all

sphingolipids, such as the major complex sphingolipids

glucosylceramides (GluCer) and glycosylphosphoinositol

ceramides (GIPCs) (Sperling and Heinz, 2003; Markham

et al., 2006).

The sphingolipid biosynthetic pathways and their cellular

compartmentalization along the secretory pathway is com-

plex (see Raffaele et al., 2009b for a review), although it has

been shown that the enzymes are mostly localized in the ER

(Marion et al., 2008; Melser et al., 2010). The first step in

ceramide biosynthesis is carried out by the enzyme complex

serine-palmitoyltransferase (Chen et al., 2006; Bach and

Faure, 2010). This enzyme is made of two subunits, LCB1

and LCB2, in charge of synthesizing the long chain bases

(LCBs, or sphingoid bases), which are specific to sphingoli-

pids. These LCBs associate with very long chain fatty

acyl-CoA (VLCFA-CoA) to form sphingolipids, which is

orchestrated by ceramide synthases. Synthesis of complex

sphingolipids such as GluCer or GIPCs is dependent on

GC synthase and IPC synthases, and constitute two different

biosynthetic pathways (Figure 1).

Previous reports on sphingolipid function in plants dem-

onstrate that interfering with their synthesis has conse-

quences for the normal functioning of the endomembrane

system (Coursol et al., 2003, 2005; Liang et al., 2003; Zheng

et al., 2005; Melser et al., 2010). However, the possible

impact of sphingolipids on the typical architecture of the

plant endomembrane system has not yet been studied in

any detail. To address these questions, we have used BY2

cells (Nagata et al., 1992) expressing different specific GFP-

tagged markers for plant cell compartments. Their short cell

cycle (11–15 h), polar ribbon-shaped organization, ease of

DNA transformation with GFP constructs and the fact that

they do not possess chloroplasts make BY2 cells extremely

suitable for these studies. We reported previously that

interfering with sterol biosynthesis may have serious

consequences for the cellular organization of BY2 cells

(Merigout et al., 2002). Therefore, it is of particular interest

to analyse the effects of sphingolipid inhibitors on plant

endomembrane organization in a similar way, and in the

same model system. The effects of fumosinin B1 (FB1, a

mycotoxin acting as a specific inhibitor of ceramide

synthase; Tsegaye et al., 2007) were analysed with the

morphodynamics and ultrastructural features of each

compartment of the secretory pathway.

Our results combine cytometry, bio-imaging and electron

microscopy approaches (Brown et al., 2007), and provide

compelling evidence for a role of sphingolipids during cell

growth and the establishment of cell polarity in higher plant

cells through their contribution to the functional organiza-

tion of the endomembrane system.

RESULTS

Fumonisin B1 affects cell growth and delays the

G2/PM phase

The effects of 1 lM FB1 were tested on 3-day-old wild-type

BY2 cells growing either in liquid cell suspension or on solid

agar medium (exponential growth phase). FB1 treatment for

Figure 1. Cell compartmentation of sphingolipid synthesis in higher plant

cells, outlining the potential pathway targeted by fumonisin B1 (FB1) and

brefeldin A (BFA). ER, endoplasmic reticulum; GC, glucosylceramide; GIPC,

glycosylinositol phosphorylceramide; LCBs, long chain bases; VLCFA, very

long chain fatty acids component.

Sphingolipids and plant membrane trafficking 959

ª 2011 The AuthorsThe Plant Journal ª 2011 Blackwell Publishing Ltd, The Plant Journal, (2011), 65, 958–971

24 h resulted in a decrease of the pack cell volume of cul-

tured cells in liquid medium by one-third compared with the

control (data not shown). Similarly, the growth of the cell

population was strongly inhibited when cells were grown

on FB1-containing solid medium (Figure 2a).

Flow cytometry analyses revealed that FB1 does not target

one phase of the cell cycle in particular (Figure 2b). After

24 h of FB1 treatment, the number of treated cells in G2/M

phase increased compared with control cells (Figure 2b,

top). The same was observed in cells that were treated with

FB1 for 72 h (Figure 2b, bottom). These results may reflect

the fact that treated cells tend to accumulate or enter the

G2/M phase more slowly (higher value and longer times).

On the other hand, 72 h of treatment with FB1 showed an

increased number of cells in the S phase (Figure 2b, bot-

tom). These results show that the growth decrease in the

BY2 cell culture corresponds to a general slowdown of the

cell cycle, possibly related to FB1 effects on the cell division

processes occurring during the M phase.

The mitotic index values in BY2 cells from 4–6-day-old

cultures (Figure 2c) confirmed that FB1 has a strong effect

on division processes. In control cells, the mitotic index

value regularly decreased over time, and at day 7, no mitosis

was recorded (not shown). In FB1-treated BY2 cells, there

was an initial strong decrease in the mitotic index value after

24 h of FB1 treatment. However, surprisingly, the mitotic

index rose again, and reached higher values than in control

cells at the same age. Light microscopy observations

confirmed that mitotic cells were still recorded after 6 and

7 days in FB1-treated cultures. These results suggest that

FB1-treated cells may be delayed in entering the cell division

processes, and would stay in the mitotic phase for a longer

period of time.

To verify whether these FB1 effects are related to effects

on BY2 sphingolipid synthesis, lipid compositions were

analysed by HPLC (Figure S1). The results showed increased

levels of DHS and 1,4-anhydro-t18:0, presumably because

these are substrates of ceramide synthases; however, the

unsaturated LCBs, which are found in complex sphingoli-

pids downstream of the ceramide synthase step, decreased.

Considering the large changes observed in LCB profile,

in particular with the accumulation of DHS as a hallmark

of ceramide synthase impairement, we assume that FB1

inhibited ceramide synthesis in BY2 cells.

Fumonisin B1 affects cell division and cell shape in a

cytoskeleton-independent manner

BY2 cells are typically organized as a multicellular ribbon

(Figure 3a). This typical morphology was strongly affected by

FB1 treatment (Figure 3b–d). After 24 h of treatment cells

tended to swell (Figure 3b), suggesting a change in the cell

growth axis. A fluorescein di-acetate test showed that the cell

viability was not affected, even after 72 h of treatment (data

not shown). The typical ribbon shape was still recognizable,

Figure 2. Effects of fumonisin B1 (FB1) on BY2 cell growth.

(a) Three-day-old BY2 cells were streaked on solid medium without (left) or with 1 lM FB1 (right). Photographs were taken after 15 days. Scale bar: 1 cm.

(b) Histograms summarizing cell cycle analysis by flow cytometry of control (white column) and FB1-treated (black column) BY2 cells (mean of three experiments).

Top: 24-h treatment (induces a significant decrease in cell population in the S and G2/M phases). Bottom: 72-h treatment results in a significant decrease of cells in

the G1 phase. Note the decrease in S phase in control cells, and a still active DNA synthesis in treated cells after 72 h of FB1 treatment. *Student’s t-test.

(c) Mitotic index of 3-day-old BY2 cells after 24, 48 and 72 h of treatment with FB1 (mean of four experiments). After 24 h of treatment, there was a significant

decrease in the mitotic index in treated cells (black column) compared with control cells (white column). Over the next few days, mitotic indices in control cells (white

column) progressively decreased, whereas in treated cells a significant level of mitotic activity remained after 48 and 72 h of treatment.

960 Anne Aubert et al.


albeit made up of enlarged, swollen cells (Figure 3c). FB1 also

induced unusual division planes, as two adjacent cells on a

radial plane were regularly observed (Figure 3d).

FM4-64 was used as a cell plate marker in order to study

possible cell plate defects in more detail (Bolte et al., 2004).

FB1-treated cells often exhibited cell plates with unusual

profiles (wavy profiles in Figure 4b,d; abnormal division

planes in Figure 4c), and a significant number of incomplete

cell plates was observed, compared with control cells

(Figure 4a). These later observations are actually in accor-

dance with the observed delay in the cell progression during

the mitotic stage described above.

Figure 3. Effects of fumonisin B1 (FB1) on BY2

cell shape and division planes, by differential

interference contrast (DIC). Note the enlarge-

ment of the cells after 24 (b) and 72 h (c) of

treatment with FB1. The typical ribbon shape of

control BY2 cells is altered (a), and the cells

exhibit abnormal division planes (d and c,

arrow). Scale bar: 10 lm.

Figure 4. FM4-64 labeling of plasma membrane

and cell plates in BY2 cells. (a) Control cells with

linearly growing cell plate. (b–d) Abnormal cell

plate patterns induced by fumonisin B1 (FB1),

exhibiting either wavy patterns (b, d), or incom-

plete/fragmented cell plates (c, d). Scale bar:

10 lm.



The cytoskeleton is known to be directly involved in these

biological processes, and it was subsequently investigated

whether FB1 altered the actin and/or microtubule networks.

The immunolabeling of actin did not reveal changes in actin

polymerization or in the actin 3D pattern (data not shown).

Similarly, the immunolabeling of microtubular arrays did

not detect changes in microtubule polymerization: the same

long tubular structures organized in a cortical network, as

well as the alignment of short structures in the phragmo-

plast on each side of the growing cell plate in dividing cells,

were observed in control and treated cells (Figure 5a–d).

However, the typical helical 3D organization of the microtu-

bules in interphase cells was disrupted (Figure 5b), and the

3D structure of the phragmoplasts in treated cells was not

as straight as in control cells (Figure 5c), reminding us of

the altered cell plate profiles shown by FM4-64 staining,

such as the wavy patterns (Figure 5d).

The loss of the typical helical pattern could relate to

changes in cell growth axis, and/or to defects in some

anchoring processes of microtubules at the plasma mem-

brane caused by the sphingolipid synthesis alterations

investigated below.

Fumonisin B1 affects the organization of the plasma

membrane

The effects of FB1 on a BY2 cell line expressing PIN1-GFP

(Boutte et al., 2006) were investigated. In control cells, PIN1-

GFP shows polar distribution, highlighting the radial mem-

branes of the cells (Figure 6a). Cell plates in dividing cells

were also visible in the PIN1-GFP line (see Boutte et al.,

2006). After treatment with FB1, the distribution of PIN1-GFP

in the cell division planes was altered (Figure 6b,c), as well

as in the growing/developing cell plate (Figure 6e). More-

over, FB1-treated cells may exhibit a loss of polar distribu-

tion in PIN-GFP, as fluorescence was often redistributed over

the whole cell surface (Figure 6d). Finally, some FB1-treated

cells exhibited intracellular PIN1-GFP-labeled aggregates,

often positioned in close proximity to the nucleus

(Figure 6f).

FB1 effects on the redistribution of PIN1-GFP polar

patterns could be related to specific alterations of secretory,

endocytic and recycling events (Boutte et al., 2006, 2007b;

Dhonukshe et al., 2007). To explore this hypothesis,

a disruption of sphingolipids in BY2 cells expressing

distinct markers for endomembrane compartments was

undertaken.

Endoplasmic reticulum and Golgi markers are affected

by FB1

The localization of GFP-HDEL, a marker for the ER, as well as

of ST-GFP, a Golgi marker, was affected by FB1 treatment

(Figure 7). Firstly, control cells expressing GFP-HDEL

showed the typical extensive fluorescent tubular network

previously described for the ER (Merigout et al., 2002). This

was mainly visible near the nuclear and cortical zone of

the cells (Figure 7a). After several hours of FB1 treatment, ER

tubules thickened and the meshwork started to fragment

(Figure 7b). At this stage, GFP labeling of the cell plate in

dividing cells was also detected, even at the end of telophase

(Figure 7d). After 24 h of treatment the ER network was GFP-

labeled, but, in addition, labeled aggregates were seen in the

vicinity of nuclear membranes (Figure 7c).

The ST-GFP labeling in the Golgi apparatus was typically

detected as numerous 1-lm fluorescent bodies distributed

over the cytoplasm (Figure 7e). The first changes were

observed after 5 h of treatment with FB1, when larger,

globular structures were detected (Figure 7f). After 24–48 h

of treatment, all the fluorescence was concentrated in

perinuclear aggregates, and nuclear membranes were also

GFP-labeled (Figure 7g). In dividing cells, GFP-labeled cell

plates were visible, as well as fluorescent nuclear

membranes in telophase cells (Figure 7h). The latter

observation was reminiscent of GFP-HDEL labeling of the

ER and nuclear membrane, and suggested that at least part

of the ST-GFP fluorescence was located or relocated to

the ER.

Figure 5. Immunofluorescence labeling of microtubules in control (a and c) or

fumonisin B1 (FB1)-treated cells (b and d). Note the altered 3D helical

organization of the microtubular pattern in interphase cells (b), and the

unusual wavy profile of the phragmoplast in dividing cells (d). Scale

bar: 10 lm.



The photoactivable ST-Kaede BY2 cell line was used to

further discriminate the dynamics of these aggregates

formation.

FB1 affects trafficking from the ER to the GA

ST-Kaede fluorescence can be converted from green to red

upon exposure to UV light (Brown et al., 2010). In control

cells, at 24 h post-photoconversion, the red (photoconvert-

ed) and green (newly synthesized) fluorescence was present

on the same Golgi stacks (Figure 8a–c). This clearly showed

that most of the newly synthesized ST-Kaede (green) is

transported to existing Golgi stacks (red), as previously

described by Brown et al. (2010). FB1 was added either 4 or

24 h before the photoconversion, in order to analyse the

effects of FB1 on ST-Kaede trafficking from ER to the Golgi.

When cells were UV-illuminated after 4 h of treatment with

FB1, no effect of FB1 was detected; at 24 h post-conversion

(i.e. after 28 h of FB1 treatment), the Golgi stacks were

labeled with both red and green fluorescence (Figure 8d and

f), suggesting that FB1 did not have severe effects on the

existing Golgi stacks, and that transport of newly-synthe-

sized ST-Kaede into the Golgi continues at a certain level.

However, green-fluorescent aggregates, like those observed

in the FB1-treated ST-GFP-expressing cells, were also

observed (compare Figure 8e with Figure 7g), and these

structures constitute the largest part of the green fluores-

cence. This indicates that most of the newly synthetized

ST-Kaede ended up in perinuclear aggregates.

When cells were UV-illuminated after 24 h of treatment

with FB1, fluorescent aggregates were already present at the

time of the photoconversion. At 24 h post-conversion (Fig-

ure 8g–i), distinct red fluorescent punctate structures are

observed, embedded in a more diffuse red fluorescent area

in the perinuclear zone (Figure 8g). These red punctate

structures, which are thought to be Golgi stacks, had to be

the Golgi stacks already present at the time of photocon-

version, and have remained resistant to FB1 treatment. The

diffuse perinuclear fluorescence contained red and green

fluorescence, reflecting accumulation of ST-Kaede in this

area both before and after photoconversion (Figure 8h–i).

This suggests that newly synthetized ST-Kaede failed to

reach the Golgi stacks, and instead accumulated in an

unknown structure around the nucleus.

As a whole, these results suggest that sphingolipid

deficiency impacts the transport from the ER to the Golgi.

Endocytosis is slightly affected by FB1

In order to further discriminate between specific FB1-

induced alterations of exocytosis versus endocytosis path-

ways, we followed the internalization of FM4-64 in BY2 cells

expressing PIN1-GFP (Figure 9a–f) and ST-GFP (Figure 9g–l).

FM4-64 internalization processes still took place under FB1

treatment (Figure 9e,k). At the same time, FB1 induces PIN1-

GFP and ST-GFP aggregates, as previously described (Fig-

ure 9d,j, respectively). Merged pictures (Figure 9f,l) show,

however, that FM4-64 did not label such aggregates. These

(a) (b) (c)

(d) (e) (f)

Figure 6. The effects of fumonisin B1 (FB1) on PIN1-GFP BY2 cell lines. Control cells (a) and FB1-treated cells (b–f), showing defects in plane division and polar

labeling (b, arrow; c, differential interference contrast, DIC, image), redistribution of PIN-GFP to longitudinal membranes (d, arrows) and unusual wavy profiles of

cell plates (e). PIN1-GFP was also observed as aggregates in perinuclear areas (f, arrow). Scale bar: 10 lm.



results suggest that in our experimental conditions and in

BY2 cells, FB1 affects the exocytosis pathways, but only

slightly effects the endocytosis pathway.

Ultrastructural analyses reveal the formation of ER-

derived membranous compartments and a transport block-

age from the ER to the GA.

Severe alterations of internal membranes and unusual

proliferations of tubular membranes were observed (com-

pare Figure 10a with Figure 10c–e). These unusual tubular

structures radiate from a clear central zone (Figure 10c,e–f),

and may correspond to the fluorescent aggregates seen

by confocal laser scanning microscopy. In contrast, the

ultrastructure of the Golgi stacks was not altered by FB1

(Figure 10b and d), even after 48 h of FB1 treatment

(Figure 10f), confirming the light microscopy observations

on ST-Kaede photoconverted cell lines. Typical ER struc-

tures were also detected in the cytoplasm (Figure 10e), and

neither nuclear or vacuolar membranes were altered by the

drug. Taken together, this suggests that the membranous

aggregates may represent newly formed ER-derived struc-

tures, named FB1-induced compartments.

We hypothesized that the altered fluorescent patterns

of the Golgi marker ST-GFP observed by light microscopy

could be related to an altered trafficking of the protein from

the ER to the GA. Immunogold labeling of ST-GFP in both

control and FB1-treated cells confirmed this hypothesis

(Figure 11a), as a significant decrease in gold labeling

on the Golgi stacks of FB1-treated cells is observed

(Figure 11b). In control cells, gold particles were rarely

seen in the cytoplasm or ER lumen, whereas in treated cells

‘clouds’ of gold labeling were detected in FB1-induced

compartments, i.e. areas of tubular membrane profusions

induced by the drug (Figure 11c,d). This demonstrates that

ST-GFP is trapped in membranous structures that are

probably ER-derived.

This labeling therefore explained the observed formation

of fluorescent aggregates after FB1 treatment labeled with

ST-GFP, ST-Kaede or PIN-GFP (Figures 6–8), i.e. proteins

(a) (b) (c) (d)

(e) (f) (g) (h)

Figure 7. The effects of fumosinin B1 (FB1) on 3D fluorescent patterns of GFP-HDEL (a–d) and ST-GFP BY2 cell lines (e–h). Scale bar: 10 lm.

(a) In control cells, GFP-HDEL labeled the endoplasmic reticulum (ER) meshwork. (b) After 5 h of treatment with FB1, the ER meshwork is still recognizable dispersed

throughout the cortical cytoplasm. (c) After 24 h of treatment with FB1, GFP-HDEL labeling is concentrated in perinuclear aggregates. Note the fluorescent nuclear

membranes. (d) After 8 h of treatment with FB1, dividing cells with a highly fluorescent cell plate wrapped in a fluorescent meshwork are visible. Labeling the nuclear

membrane.

(e) In the control cell, ST-GFP labeled the Golgi stacks, giving a punctate pattern throughout the cytoplasm. (f) After 5 h of treatment with FB1, a thickening of

fluorescent units is observed. (g) The ST-GFP fluorescence is concentrated in perinuclear aggregates and nuclear membranes. These features are somehow similar

to the ones described in the 24-h treatment of GFP-HDEL cell lines (compare with Figure 6c). (h) After 24 h of treatment with FB1, dividing cells exhibit a fluorescent

cell plate, with fluorescent aggregates at the cell plate extremities; there is strong labeling of the nuclear membrane.



on their way to anterograde transport being blocked in

ER-derived structures. The fact that no visible difference in

ultrastructural features of plasma membrane or endocytic

compartments was detected also confirmed our previous

hypothesis that endocytosis dynamics were not primarily

affected by the drug.

FB1-induced compartments are distinct from BFA-induced

reorganization of the ER–GA complex

The next question was to know how specific the FB1-induced

modifications were compared with some other ER–GA-

disrupting drug, such as BFA (Satiat-Jeunemaitre et al.,

1996). After treatment with BFA, the Golgi stacks of BY2 cells

underwent profound reorganization, leading to the forma-

tion of ER–Golgi hybrid compartments (Figure 12c;

Langhans and Robinson, 2007), and the typical membrane

profusions induced by FB1 treatment (Figure 12b) were

never observed. Interestingly, when FB1-treated cells were

treated with BFA for 1 h, the cells exhibited the two types of

abnormal figures: ER/Golgi hybrid compartments typical of

BFA treatment; and ‘FB1-induced compartments’ made

of a profusion of tubular and swollen membranes (compare

Figure 12a and d). These results outline a specific sensitivity

to each drug, confirming fully distinct molecular targets.

They also confirm that the formation of FB1-induced

compartments can be regarded as a ‘de novo’ ER-derived

(a) (b) (c)

(d) (e) (f)

(g) (h) (i)

Figure 8. The effects of fumosinin B1 (FB1) on the 3D fluorescent pattern of ST-Kaede. Cells were treated for short (4 h) or long (24 h) periods with FB1 (d–h and g–i,

respectively), were photoconverted from green to red and then observed 24 h later. In this plate, the fluorescence detected in the red channel (a, d, g) is therefore

associated with ST-Kaede proteins in place before photoconversion, meanwhile the green fluorescence (b, e, h) is associated with newly synthesized ST-Kaede

during the 24 h period post-photoconversion. Scale bar: 10 lm.

(a–c) Control ST-Kaede cells. (a) Red ST-Kaede Golgi stacks in place at the time of photoconversion. (b) Newly synthesized green ST-Kaede labeling of Golgi stacks.

(c) The merged picture suggests that most of the newly synthesized Kaede proteins have reached pre-existing Golgi compartments.

(d–f) Treatment with FB1 for 4 h. (d) Red ST-Kaede labeling of Golgi stacks in place at the time of photoconversion. (e) Newly synthesized ST-Kaede proteins

accumulate in fluorescent aggregates and form punctuate structures. (f) Merged pictures: newly synthesized proteins have reached pre-existing Golgi stacks or have

accumulated in non-Golgi structures.

(g–i) After 24 h of treatment with FB1. (g) Red ST-Kaede labeling of Golgi stacks in place at the time of photoconversion. (h) Newly synthesized ST-Kaede proteins

accumulate in fluorescent aggregates; they do not form punctate structures. (i) Merge pictures: newly synthetized proteins accumulate in non-Golgi structures.



compartment, rather than a reorganization of previous

compartments, as found in BFA-treated cells.

DISCUSSION

These results show that, in BY2 cells, an FB1-sensitive

sphingolipid pathway impairs several cell functions, inter-

fering with the membrane trafficking pathways by: (i)

introducing a delay in the progression through the cell cycle;

(ii) introducing defects in cell plane positioning, cell plate

growth and polar axis establishment; (iii) inducing the

ER-derived membranous tubular network, i.e. FB1-induced

compartments; and (iv) blocking the transport of cargo

molecules from the ER to the GA.

FB1 effects outline a low sphingolipid turnover and

a cell ability to compensate sphingolipid deficiency

The peculiarity of FB1 treatment compared with other

pharmacological agents disturbing the organization of cell

(a) (b) (c)

(d) (e) (f)

(g) (h) (i)

(j) (k) (l)

Figure 9. Effects of fumosinin B1 (FB1) on FM4-64 internalisation in PIN1-GFP (a–f) and ST-GFP BY2 (g–l) cells.

(a–c) PIN1-GFP control cell: (a) typical fluorescent PIN1 pattern; (b) internalization of FM4-64; (c) the merged picture shows the partial localization of FM4-64 and GFP

fluorescence. Scale bar: 10 lm.

(d–f) PIN1-GFP-treated cells: (d) FB1-induced perinuclear aggregates; (e) internalization of FM4-64; (f) the merged picture shows no co-localization of FM4-64 with

FB1-induced aggregates. Scale bar: 10 lm.

(g–i) ST-GFP control cells: (g) typical fluorescent Golgi pattern; (h) internalization of FM4-64; (i) the merged picture shows the partial co-localization of FM4-64 with

GFP-labeled organelles. Scale bar: 10 lm.

(j–l) ST-GFP-treated cells: (j) FB1-induced aggregates; (k) internalization of FM4-64; (l) no co-localization of FM4-64 with ST-GFP aggregates. Scale bar: 10 lm.



membranes, such as BFA (Satiat-Jeunemaitre and Hawes,

1992; Satiat-Jeunemaitre et al., 1996; Merigout et al., 2002;

Ritzenthaler et al., 2002; Couchy et al., 2003; Langhans

and Robinson, 2007) or nordihydroguiaretic acid (NDGA)

(Merigout et al., 2002), is the time period needed to detect

any FB1 effects on cell growth. The low turnover of sphin-

golipids could explain these slow changes in subcellular and

cellular morphological features. On the other hand,

biochemical changes have been reported to occur long

before any morphological changes are observed (Abbas

et al., 1994; Spassieva et al., 2002). This may indicate that

plants are able to adjust their growth mechanisms to

compensate for a reduction in ceramide synthesis, a process

already noticed in the case of the partial suppression of the

early step of sphingolipid synthesis (Chen et al., 2006).

Impact of sphingolipids deficiency on cell polarity

A deficiency in sphingolipids has a strong impact on the

establishment of a polar axis, and may lead to the mislo-

calization of the PIN1 proteins. Lateral diffusion of PIN

labeling, together with a disorganization of the microtubular

array, over the cell surface has previously been described

when BY2 cells were turned into protoplast (Boutte et al.,

2006). In this study, the observations suggest that the

(a) (b)

(c) (d)

(e) (f)

Figure 10. Effects of fumosinin B1 (FB1) on the cell ultrastructure. (a) Control with typical plant cell endomembrane features. (b) Golgi stack in control cells.

(c, d) Micrographs of cell ultrastructure after 24 h of treatment with FB1 at different electron microscope magnifications. Golgi stack and endoplasmic reticulum (ER)

tubules are recognizable: nuclear and vacuolar membranes appeared unaffected, and the Golgi stack morphology is also unaffected.There is an extensive tubular

membrane network (tn) within the cytoplasm.

(e, f) Higher magnifications often outline the peculiar radiating pattern of this network from a 0.5–1 lm square electron-transparent area (*).

Key: er, endoplasmic reticulum; ga, Golgi apparatus; n, nucleus; v, vacuole.

Scale bars: a, c, e, f, 1 lm; b, d, 200 nm.



mechanisms involved in the maintenance of cell polarity

could be related to the fine organization of the plasma

membrane. In our study, such effects could indeed be

related to a profound alteration of plasma membrane orga-

nization. PIN1 proteins have been reported to be associated

with lipid rafts in plants (Titapiwatanakun et al., 2009), and

lipid rafts are known to be enriched in sterols and sphingo-

lipids (Mongrand et al., 2004; Raffaele et al., 2009a). More-

over, proteins and sphingolipids associate to form

microdomains in the trans-Golgi network (Barz and Walter,

1999). Therefore, the redistribution of PIN1 proteins may

be related either to a change in raft composition or to a

defect in protein sorting along the secretory pathway.

Sphingolipids are involved in the orchestration

of anterograde flows

Our results support the hypothesis that sphingolipids are

essential to maintain anterograde flows, as ceramide defi-

ciency causes an inhibition of ER to GA transport of cargo

molecules. Such effects of sphingolipid deficit were also

seen in animal cells where it was shown that transport of GPI

proteins from ER to the GA was dependent on sphingolipids

(Barz and Walter, 1999). Formations of abnormal cell plate or

defects in cell plate growth under FB1 are also good indi-

cators for involvement of sphingolipids in the regulation of

exocytosis. The fundamental question of how this blockage

is related to changes in ultrastructural organization remains.

In FB1 treatment, the deficiency of sphingolipids and

alterations of subcellular processes are associated with a

profusion of ER-derived tubular membrane networks. As

FB1 may induce apoptosis-like cell death in plants (Asai

et al., 2000; Shi et al., 2007), such an occurrence of mem-

branes could be related to the formation of apoptotic-like

bodies along the secretory pathway. These modifications

are fully distinct from the ones induced by the ER-to-GA

transport inhibitor BFA, where Golgi membranes and ER

membranes fuse and create a hybrid structure (Langhans

and Robinson, 2007). FB1 did not change the endomem-

brane reactivity to BFA: as these ER-derived structures are

full of newly synthesized cargo molecules, they suggest that,

without sphingolipids, ER membranes are not able to favor

vesicle shuttle between the ER and the GA, but favor some

tubular expansions.

As a whole, this study demonstrates specific roles for

sphingolipids in endomembrane organization. Therefore.

the availability of the sphingolipid pool could be a limiting

factor in the ‘differentiation’ of ER membranes towards other

compartments (Figure 13). Modification of the sphingolipid

(a) (c)

(b) (d)

Figure 11. Immunogold labeling of Golgi stacks in ST-GFP cell lines by anti-GFP antibodies (cryofixation by high-pressure freezing). Scale bar: 200 nm.

(a) Control cells.

(b) Quantification of gold particles on 60 Golgi stacks in control cells (white column) or in fumosinin B1 (FB1)-treated cells. Note the strong decrease of

immunolabeling in treated cells.

(c, d) Immunolabeling of the FB1-induced tubular meshwork by anti-GFP, illustrating the trapping of ST-GFP molecules within the area. Scale bar: 500 nm.



molecular signature would therefore interfere with specific

membrane protein-associated molecular machinery in

charge of vesicular mobility or fusions that could character-

ize ER-to-GA traffic or GA morphogenesis.

EXPERIMENTAL PROCEDURES

Biological material

Nicotiana tabacum cv. Bright Yellow 2 (BY2) suspension-culturedcells were grown in modified MS medium as previously described(Couchy et al., 2003). Four BY2 transgenic cell lines were used in thisstudy: ST-GFP as a Golgi complex marker (Saint-Jore et al., 2002);ST-Kaede (Brown et al., 2010); GFP-HDEL as an ER marker (Merigoutet al., 2002); and PIN1-GFP as a cell polarity marker Boutte et al.,2006). Experiments were performed on 3-day-old BY2 culturesexhibiting a high mitotic index.

Drug treatments

Fumosinin B1 (FB1 F1147; Sigma-Aldrich, http://www.sigmaaldrich.com) was added to the suspension culture from a 10 mM aqueousstock solution, to reach a 1 lM final concentration.

Treatments with BFA in a final concentration of 10 lg ml)1

(Sigma-Aldrich) were performed for 1 h as previously described(Merigout et al., 2002).

Cytological, immunocytological approaches and probes

for light microscopy

DNA was stained with Hoechst 33342 (2 lg ml)1). FM4-64 dye wasadded to the cell suspension as described in Bolte et al. (2004).

Indirect immunofluorescence experiments were performed aspreviously described (Hawes and Satiat-Jeunemaitre, 2001).Anti-tubulin antibodies (Interchim, http://www.interchim.com) wereused at a dilution of 1:100. An Alexa Fluor 488-conjugated anti-mouse antibody (Invitrogen, http://www.invitrogen.com) was usedas a secondary antibody at a dilution of 1:400.

Microscopy

Imaging by light microscopy. Cells expressing GFP or Kaedefusions were imaged using confocal microscopy (Leica SP2; Leica,http://www.leica.com). Single and dual color imaging were per-formed as previously described (Brown et al., 2010). Transmissionimages were taken simultaneously in Nomarski mode differentialinterference contrast (DIC). UV-violet light exposure was used forthe photoconversion of Kaede protein.

Electron microscopy

The BY2 cells were processed as described in Hawes and Satiat-Jeunemaitre (2001), except that the osmium post-fixation step wasreplaced by a mixture of 1% osmium and 1.5% potassium ferrocy-anide. Specimens were embedded in epoxy resin (Agar low-viscosity premix kit medium, Oxford Instruments, Saclay, France)and polymerized for 16 h at 60�C. Selected pictures are represen-tative of observations performed on 20 sections resulting from threeexperiments. Alternatively, embedding through an automativemicrowave tissue processor (AMW) device (Leica) was used, fol-lowing the manufacturer’s instructions. For immunogold labeling,BY2 cells were high-pressure frozen (EMPACT2; Leica) and freezesubstituted (AFS2; Leica). Specimens were then infiltrated andembedded in LRWhite resin. Immunogold labeling was performed

(a) (b)

(c) (d)

Figure 12. Brefeldin A (BFA) and fumonisin B1 (FB1) have distinct effects on the endomembrane network. (a) Control cell. (b) FB1-treated cells with typical tubular

membrane profusion. (c) BFA-treated cell with endoplasmic reticulum (ER)–Golgi apparatus (GA) hybrid structure. (d) FB1- and BFA-treated cell showing both ER–

GA hybrid structures (arrow) and membrane profusion (arrowhead). Key: er, endoplasmic reticulum; ga, Golgi apparatus; tn, tubular membrane network; v, vacuole.

*Electron-transparent area. Scale bar: 200 nm.



using rabbit GFP antibodies (AB290; Abcam, http://www.abcam.com), at a dilution of 1:400.

Ultrathin sections (70–90 nm, Ultracut UC6; Leica) were post-stained with aqueous 2% uranyl acetate/lead citrate, as described byHawes and Satiat-Jeunemaitre (2001). Grids were examined under aJEOL 1400 TEM operating at 120 kV (JEOL, http://www.jeol.com).Images were acquired using a post-column high-resolution(11 megapixels) high-speed camera (SC1000 Orius; Gatan, http://www.gatan.com).

Flow cytometry

Cell cycle analysis of isolated nuclei. Isolated nuclei from a2–5-day-old cell culture were stained with propidium iodide (IP;50 lg ml)1) (Coba de la Pena and Brown, 2001), and data wereacquired (excitation at 532 nm; emission through a 590-nm long-pass filter) on a CyFlow SL cytometer (Partec SL, http://www.partec.com). Cell cycles were analysed using MultiCycleAV software(P. Rabinovitch, University of Washington).

Lipid analysis

The analysis of total long chain bases of sphingolipids was per-formed by HPLC after fluorescent derivatization, as described before(Bach et al., 2008).

Statistical analysis

Data are means � SEs of a minimum of three independent experi-ments. Differences between means were evaluated by Student’s

t-test with P < 0.05 being taken as the level of significance(*P < 0.05).

ACKNOWLEDGEMENTS

The BY2 cell lines expressing ST-GFP were kindly provided byC. Hawes (Oxford, UK). We thank Marie Noelle Soler and SpencerBrown for their help in imaging and flow cytometry approaches.Anaıs Carpentier established the protocol for AMW processing ofplant cells. Thanks are also due to Cynthia Dupas and Karim Hdidoufor maintenance of the BY2 culture. BSJ gratefully acknowledgesfunding from the IFR87 and from the ANR ‘Sphingopolar’ (ANR-07-BLANC-0202). SA was financed by the EC Lifelong Learning pro-gram. This work used the facilities of the cell biology unit of theImagif platform of the Centre de recherche de Gif sur Yvette (http//http://www.imagif.cnrs.fr), supported by the Conseil General del’Essonne.

SUPPORTING INFORMATION

Additional Supporting Information may be found in the onlineversion of this article:Figure S1. The effects of fumonisin B1 (FB1) on total long chain basecontents in BY2 cells.Please note: As a service to our authors and readers, this journalprovides supporting information supplied by the authors. Suchmaterials are peer-reviewed and may be re-organized for onlinedelivery, but are not copy-edited or typeset. Technical supportissues arising from supporting information (other than missingfiles) should be addressed to the authors.

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Sphingolipids involvement in plant endomembrane differentiation: the BY2 case