Transient transformation and RNA silencing in Zinnia tracheary element differentiating cell cultures


  • Satoshi Endo,

    1. RIKEN Plant Science Center, Yokohama, Kanagawa 230-0045 Japan
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    • Present address: Umeå Plant Science Centre, 901 83 Umeå, Sweden.

  • Edouard Pesquet,

    1. UMR-CNRS-UPS 5546 «Surfaces Cellulaires et Signalisation chez les Vegetaux», Pole de Biotechnologie Vegetale, 24 Chemin de Borde-Rouge, 31326 Castanet-Tolosan, France
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    • Present address: Umeå Plant Science Centre, 901 83 Umeå, Sweden.

  • Gen Tashiro,

    1. RIKEN Plant Science Center, Yokohama, Kanagawa 230-0045 Japan
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  • Hideo Kuriyama,

    1. RIKEN Plant Science Center, Yokohama, Kanagawa 230-0045 Japan
    2. Department of Biological Sciences, Graduate School of Science, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033 Japan
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  • Deborah Goffner,

    1. UMR-CNRS-UPS 5546 «Surfaces Cellulaires et Signalisation chez les Vegetaux», Pole de Biotechnologie Vegetale, 24 Chemin de Borde-Rouge, 31326 Castanet-Tolosan, France
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  • Hiroo Fukuda,

    1. Department of Biological Sciences, Graduate School of Science, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033 Japan
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  • Taku Demura

    Corresponding author
    1. RIKEN Plant Science Center, Yokohama, Kanagawa 230-0045 Japan
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This article is corrected by:

  1. Errata: Correction Volume 54, Issue 5, 964, Article first published online: 12 May 2008

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The Zinnia elegans cell culture system is a robust and physiologically relevant in vitro system for the study of xylem formation. Freshly isolated mesophyll cells of Zinnia can be hormonally induced to semisynchronously transdifferentiate into tracheary elements (TEs). Although the system has proven to be valuable, its utility is diminished by the lack of an efficient transformation protocol. We herein present a novel method to introduce DNA/RNA efficiently into Zinnia cells by electroporation-based transient transformation. Using reporter gene plasmids, we optimized the system for efficiency of transformation and ability for the transformed cells to transdifferentiate into TEs. Optimal conditions included a partial digestion of the cell walls by pectolyase, a low voltage and high capacitance electrical pulse and an optimal medium to maintain cell viability during transformation. Beyond the simple expression of a reporter protein in Zinnia cells, we extended our protocol to subcellular protein targeting, simultaneous co-expression of several reporter proteins and promoter-activity monitoring during TE differentiation. Most importantly, we tested the system for double-stranded RNA (dsRNA)-induced RNA silencing. By introducing in vitro-synthesized dsRNAs, we were able to phenocopy the Arabidopsis cellulose synthase (CesA) mutants that had defects in secondary cell-wall synthesis. Suppressing the expression ofZinnia CesA homologues resulted in an increase of abnormal TEs with aberrant secondary walls. Our electroporation-based transient transformation protocol provides the suite of tools long required for functional analysis and developmental studies at single cell levels.


Genomic data is accumulating at an ever-accelerating rate. As a result, there needs to be acceleration in the development of techniques to capitalize more efficiently on this data. One pressing need is for a rapid transformation protocol to study gene function in specific developmental processes. Currently, the main approaches used are stably transformed whole plants and/or protoplast and cell cultures. However, when using transgenic whole plants or tagged mutants in reverse-genetic approaches, phenotypic analysis can be made more difficult through a complication of primary and secondary effects of the misregulation of the gene of interest. The main problem with protoplast and cell cultures is that they generally lack a developmental context.

The Zinnia elegans cell culture system is one of the few cell culture systems exhibiting a defined developmental process (Fukuda and Komamine, 1980). In this system, freshly isolated mesophyll cells are hormonally induced to transdifferentiate into tracheary elements (TEs). This in vitro system has been used extensively since the 1980s because of its high reproducibility and efficiency (3 days to reach around 30–60% of TEs), its semisynchrony (each major developmental process occurs at the same time), its strict hormonal induction (under the combined trigger of auxin and cytokinin) and its high similarity to TE formation in planta (Milioni et al., 2002; Pesquet et al., 2003). During Zinnia TE differentiation, cells successively undergo a shift in their cytoskeleton, secondary cellulosic wall formation, programmed cell death and finally lignification (Fukuda, 1996; Kuriyama and Fukuda, 2001). In planta, TEs interconnect to constitute a network of vessels that transports water and minerals throughout the plant. The Zinnia system has been extensively used by many teams around the world to study cell cycles, cell expansion, cell-wall construction, gene regulation, programmed cell death, lignin deposition and cell-to-cell interaction mechanisms.

To date, over 18 000 expressed sequence tags (ESTs) have been sequenced from TE-differentiating cell cultures and have been analysed by microarray (Demura et al., 2002; Milioni et al., 2002; Pesquet et al., 2005), providing fundamental information for the study of gene function. However, direct modulation of the gene expression in Zinnia is currently limited because of its recalcitrance to stable transformation. Although particle bombardment-based transient transformation has been successfully used in Zinnia cultured cells (Ito and Fukuda, 2002; Nakanomyo et al., 2002), it has only been effective at relatively late culture stage (just before morphogenesis, including secondary wall formation and programmed cell death), which greatly limits its usefulness.

In this study, we have developed an electroporation-based method to introduce plasmid DNA into freshly isolated Zinnia mesophyll cells transiently without preventing TE differentiation. Beyond the simple introduction of plasmid DNA into Zinnia cells, we adapted this method for: (i) protein subcellular localization in differentiating Zinnia cells, (ii) co-expression of several reporter genes, (iii) promoter activity monitoring and (iv) RNA silencing by the direct introduction of double-stranded RNAs (dsRNAs). This allowed us to undertake functional analyses of Zinnia genes at any stage of TE differentiation.

The most important aspect of this study is the practical use of dsRNA-mediated RNA silencing. To ensure the efficiency and accuracy of the RNA silencing, dsRNA sequences corresponding to CesA genes were introduced into Zinnia cells. CesA genes have been extensively characterized in Arabidopsis and other species (Delmer, 1999; Kimura and Kondo, 2002; Taylor et al., 1999, 2000, 2003; Turner and Somerville, 1997). The Arabidopsis genes AtCesA1, AtCesA3 and AtCesA6 participate in primary wall formation (Saxena and Brown, 2005), whereas AtCesA8/IRX1, AtCesA7/IRX3 and AtCesA4/IRX5 participate in secondary cell-wall formation (Taylor et al., 2003). In this study, we succeeded in phenocopying the cellular phenotype of the irx mutants using dsRNA-mediated RNA silencing of Zinnia secondary cell-wall CesA homologues.


Introduction of plasmid DNA into freshly isolated Zinnia cells by electroporation

Electroporation conditions were first established in order to obtain the maximal transformation efficiency without affecting the capacity of the cell cultures to transdifferentiate into TEs. Transformation efficiency was monitored by introducing plasmid DNA carrying a CaMV 35S promoter fused to the green fluorescent protein (GFP) reporter gene (P35S::GFP), and by scoring the number of GFP-expressing cells. Several parameters that had the potential to affect transformation efficiency were tested (Table 1). These included: (i) the pre-treatment of cells with cell-wall hydrolytic enzymes to reduce the cell-wall barrier to transformation, (ii) the optimal medium for electropulsation (medium with HEPES and/or CaCl2), (iii) the intensity of the electric field and the salts affecting the electrical pulse, (iv) the effect of polyethylene glycol (PEG) or salmon genomic DNA as carrier DNA to increase cell transformation and (v) the maintenance of extracellular osmolarity with mannitol to ensure the highest cell viability throughout the procedure.

Table 1.   Optimization of the electroporation conditions for the highest transformation efficiency and unaltered tracheary element (TE) differentiation
StepsFactorsGFP-expressing cells (%)TE differentiationNo. observations
  1. Optimal conditions, shown in bold letters, are a 30-min pre-treatment with 0.1 mg ml−1 of pectolyase Y-23; washing in a 260 mm mannitol, 5 mm CaCl2, 5 mm HEPES, pH 7.2, solution; an electropulsation in the presence of 0.02 m KCl, 5% polyethylene glycol (PEG); and a post-culture in Fukuda–Komamine medium supplemented with 250 mm of mannitol. GFP-expressing cells were observed at 48 h of culture. TE differentiation was investigated at 72 and 96 h of culture.

  2. No effect, no GFP-expressing cell; not affected, TE differentiation occurred as that in the optimal conditions; delayed, rates of TEs were <3% at 72 h of culture; inhibited, no TE at 96 h of culture; ND, not determined.

  3. aGFP-expressing cells were observed.

Pre-treatmentPectolyase Y-230.1 mg ml−1>1Not affected>3
  0.5 mg ml−1NDaDelayed2
  1.0 mg ml−1NDaInhibited2
 Macerozyme R101.0 mg ml−1No effectInhibited2
  5.0 mg ml−1No effectInhibited2
 Macerozyme R2001.0 mg ml−1No effectInhibited2
  5.0 mg ml−1No effectInhibited2
 Cellulase Onozuka1.0 mg ml−1No effectND2
 Papain0.1 mg ml−1No effectND2
 CaCl20.2 mNo effectND1
 EDTA0.2 mNo effectND1
WashingHEPES and mannitol <1Delayed>3
  With CaCl2>1Not affected>3
 FK medium <1Not affected2
Electropulsation0.02 m KCl <1Not affected>3
  With 5% PEG>1Not affected>3
  With carrier DNA>1ND2
 0.02 m NaCl >1Inhibited>3
CultureFK mediumWith 200 mm mannitol<1Delayed>3
  With 250 mm mannitol>1Not affected>3

A hydrolytic enzyme pre-treatment was necessary because unpretreated cells were rarely transformed (data not shown). Freshly isolated mesophyll cells were treated with hydrolytic enzymes to mildly digest the primary walls. Different hydrolytic enzymes (cellulase, pectolyase, macerozymes and papain) and chemicals (EDTA and CaCl2) were tested, but only the pectolyase pre-treatment at the low concentration significantly improved transformation efficiency without inhibiting TE differentiation (Table 1).

The cells were washed in 260 mm mannitol solutions supplemented with 5 mm HEPES and/or 5 mm CaCl2. Extracellular hyperosmolarity was maintained with the higher concentration of mannitol solutions throughout the procedure, to reduce cell mortality and thereby allow a high frequency of TE differentiation (Table 1).

Electropulsation in the presence of KCl and PEG, at low intensity (325 V cm−1) and high capacitance (900 μF), giving a time constant of 45–60 msec, ensured a high transformation efficiency without affecting TE differentiation (Tables 1 and 2). NaCl prevented TE differentiation under the same electrical conditions (Table 1), and a higher electric field increased transformation efficiency but prevented TE differentiation (data not shown). The addition of carrier DNA also increased the transformation efficiency (Table 1). However, in all the subsequent experiments we omitted the carrier DNA in order to avoid the introduction of unknown genetic contaminants.

Table 2.   Effect of the electrical parameters on transformation efficiency and tracheary element (TE) differentiation
Voltage (V)Electric field (V cm−1)Capacitance (μF)Time constant (msec)GFP-expressing cells (%)TE differentiationElectrical energy (J)No. observations
  1. Cells were electropulsated in a solution with final concentrations of 0.2 m mannitol, 4 mm HEPES, 4 mm CaCl2, 0.02 m KCl, 5% polyethylene glycol (PEG) and 100 μg ml−1 plasmid DNA, with or without 25 μg ml−1 double-stranded RNA (dsRNA) at the indicated parameters. Optimal electropulsation, shown in bold letters, is a 325-V cm−1 and 900-μF electric pulse. GFP-expressing cells were observed at 48 h of culture. TE differentiation was investigated at 72 h of culture. Definitions: not affected, TE differentiation occurred as that in the optimal electropulsation; delayed, rates of TEs were <3% at 72 h of culture. Stored electrical energy is calculated as 0.5 C V02 (C, capacitance; V0, initial voltage) for each treatment.

12431060035<0.5Not affected4.61
12030080058–59>1Not affected5.8>3
13032590045–60>1Not affected7.6>3
14035070050–53>1Not affected6.9>3
15037570050–54>1Not affected7.92
15037530018–19>0.5Not affected3.4>3
15037520012–18>0.5Not affected2.3>3
200500150 9>1Not affected3.01
20050075 4<0.5Not affected1.51

Under the optimal transformation conditions [30-min pre-treatment with 0.1 mg ml−1 of pectolyase Y-23; washing in a solution of 260 mm mannitol, 5 mm CaCl2, 5 mm HEPES, pH 7.2; electropulsation at 325 V cm−1 and 900 μF in the presence of 0.02 m KCl, 5% PEG; and post-culture in Fukuda–Komamine (FK) medium (Fukuda and Komamine, 1980) supplemented with 250 mm mannitol], GFP-expressing cells totalled 1% of the cultured cells at 48 h after the electric pulse (Figure 1c). The overall transformation efficiency was apparently >1%, because the use of the GUS reporter gene gave at least 10 times as many GUS-expressing cells as GFP-expressing cells (data not shown). Transformation under the optimal conditions had no effect on TE differentiation efficiency, but delayed the TE differentiation processes by approximately 24 h (Figure 1a). Direct observation of cells after electroporation showed that palisade mesophyll cells expressed GFP (Figure 1d,e). By monitoring the number of cells with septa we confirmed that electropulsation had no affect on cell division (Figure 1b), and that GFP was visible in both undivided and dividing cells. Transformed cells successfully differentiated into TEs (Figure 1f,g). About 8% of all GFP-expressing cells were TEs, thus representing 0.08% of the total number of cells (Figure 1c). This value might be underestimated because of the programmed cell death process, which starts when TEs begin to become visible and removes all cytoplasmic contents of the cell. On the practical side, this efficiency is sufficient for further analysis because of both the high cell density and the high TE differentiation efficiency of the Zinnia system.

Figure 1.

 Time course of green fluorescent protein (GFP) expression and tracheary element (TE) differentiation after electroporation. (a) Rates of TEs in total cells: □, native culture conditions; ○, unpulsed cultures; •, pulsed cultures.
(b) Rate of cell division, estimated by the number of cells with septa, in pulsed (•) and unpulsed (○) cultures.
(c) Rates of GFP-expressing cells in pulsed (•) and unpulsed (○) cultures, shown with the scale on the left y-axis; bsl00001, with the scale on the right y-axis shows rates of GFP-expressing TEs in total GFP-expressing cells. Each data point presents the mean of three independent cultures ± SD.
(d, e) Zinnia cells electroporated with P35S::GFP plasmid and cultured for 24 h, which were observed in bright field (d) and GFP fluorescence (e).
(f, g) Differentiating TEs electroporated with P35S::GFP plasmid and cultured for 84 h, which were observed in bright field (f) and GFP fluorescence (g). Bars = 25 μm.

Many cell culture systems are often used for the subcellular localization of proteins. We carried out differential targeting assays with fluorescent proteins in Zinnia cell culture under optimal transformation conditions. Normal yellow fluorescent protein (YFP) accumulates mostly in the cytoplasm (Figure 2a), similarly to the GFP used in the optimization assays (Figure 1g). Adding a nuclear localization signal to YFP restricted its accumulation in the nucleus, thereby verifying that this system can be efficiently used for protein subcellular localization (Figure 2b).

Figure 2.

 Subcellular targeting and co-expression in differentiating Zinnia cells.
(a, b) Localization of normal yellow fluorescent protein (YFP) in differentiating tracheary elements (TEs) (a), and nuclear targeting of YFP (b).
(c–f) Co-introduction of P35S::YFP (c, e) and P35S::CFP (d, f) into isolated Zinnia mesophyll cells. YFP and cyan fluorescent protein (CFP) signals were observed in non-TE cells (c, d) and differentiating TEs (e, f). Scale bars = 25 μm.

Another common use of cell culture systems is the study of the co-expression of two or more labelled proteins. We successfully co-transformed and co-expressed YFP and cyan fluorescent protein (CFP) in Zinnia cell cultures. Both fluorescent proteins were observed not only in non-TE cells (Figure 2c,d), but also in TEs (Figure 2e,f). The co-expression efficiency was high, as about 80% of YFP-expressing cells also showed CFP expression.

We examined whether this system can be used for monitoring promoter activity during TE differentiation. The promoter activity of an early TE differentiation marker, TRACHEARY ELEMENT DIFFERENTIATION-RELATED3 (TED3), was followed during TE differentiation using a TED3 promoter::nuclear-targeted YFP (PTED3::YFPnuc) construct. It was shown that TED3 transcripts accumulated before TE morphogenesis became visible, and that the expression was maintained after TE cell death in the Zinnia system (Demura and Fukuda, 1994). As expected, YFP-expressing cells were observed through TE differentiation (Figure S1). Concerning cell specificity, the expression was observed only in some cells that did not show visible secondary cell walls. Therefore, our protocol can be used efficiently to monitor promoter activity throughout a culture at the single-cell level.

Suppression of transient expression of introduced genes by corresponding dsRNAs

In order to determine if the electroporation-based method described above could be applied to RNA silencing, two species of dsRNA were co-introduced with P35S::GFP into Zinnia cells. One of the dsRNA sequences corresponded to 0.4 kbp of the CaMV 35S promoter, and the other sequence corresponded to 0.4 kbp of the GFP sequence. dsRNAs were synthesized in vitro directly from PCR fragments amplified with RNA transcription initiation sites at both ends, which were present in the subcloning vector. Comparison of GFP expression with cells electroporated with only the P35S::GFP plasmid showed that P35S dsRNA had no significant effect on GFP expression, whereas GFP dsRNA suppressed GFP expression throughout cell culture (Figure 3a). Similarly, the reduction in GUS activity derived from P35S::GUS was specifically caused by GUS dsRNA targeted to the GUS coding sequence (Figure 3b). Therefore, introduction of dsRNAs by electroporation is effective in targeted gene silencing in the Zinnia system.

Figure 3.

 Effect of double-stranded RNA (dsRNA) on transient expression of exogenous genes in Zinnia cells. The plasmids indicated (100 μg ml−1) and/or dsRNAs (25 μg ml−1) were co-electroporated into isolated Zinnia mesophyll cells.
(a) Effects of dsRNA for the GFP open reading frame (ORF) (•) and the P35S promoter region (○) on transient GFP expression derived from P35S::GFP during Zinnia cell cultures; □, control without dsRNA. Each data point represents the mean of three independent cultures ± SD.
(b) Detection of GUS activity derived from P35S::GUS by X-Gluc staining and fluorometric assay of cells cultured for 48 h after electroporation with indicated plasmids and dsRNAs. Lambda dsRNA that corresponded to a partial sequence of lambda DNA was used as a control.

Effects of dsRNAs corresponding to CesA genes on TE morphogenesis

One of the most powerful applications for this method is the dsRNA-mediated suppression of endogenous genes functioning in TE differentiation. Three Zinnia CesA cDNA clones, Z2657, Z632 and Z811, were chosen (Demura et al., 2002; Figures S2 and S3) to test the efficacy of the technique. The purpose of using CesAs was to phenocopy, at the cellular level, the secondary cell-wall changes observed in the Arabidopsis CesA mutants (Taylor et al., 1999, 2000, 2003; Turner and Somerville, 1997).

Z2657 is identical to ZeCesA1 (Haigler et al., 2001) and is most likely to be homologous to AtCesA8 (Figures S2 and S3). The mutant allele of AtCesA8, irx1, shows defects in secondary wall formation (Taylor et al., 2000). Microarray analysis showed that Z2657 transcripts are highly expressed during the formation of specialized secondary cell walls in TEs, shown by calcofluor staining (Figure S4; Demura et al., 2002). Introduction of Z2657 dsRNA in cultured mesophyll cells resulted in an increase of TEs with abnormal secondary cell walls, compared with cells electroporated with lambda dsRNA or without dsRNA (Figure 4), whereas no major cell-wall alterations were visible in non-TE cells (data not shown). The abnormal TEs observed in the controls were probably a result of the pre-treatment of mesophyll cells by pectolyase at the beginning of culture (Figure 4). Abnormal TEs were further subdivided into four categories exhibiting defects in secondary cell walls, such as thin walls, separation of the secondary cell walls from the primary cell walls, interrupted walls and loosely coiled walls (Figure 4a–e). Based on their appearances under light microscopy, introduction of Z2657 dsRNA resulted in an increase in TEs with secondary cell walls that looked ‘thin’ and ‘separated’ (where the secondary cell wall is separated from the primary cell wall) (Figure 4g).

Figure 4.

 Effect of Z2657 double-stranded RNA (dsRNA) on secondary cell-wall formation in tracheary element (TE) differentiation.
(a–e) Abnormal TEs were subdivided into four categories exhibiting defects in secondary cell walls: thin walls (b), separation of the secondary cell walls from the primary cell walls (c), interrupted walls (d) and loosely coiled walls (e). Scale bars = 25 μm.
(f) Rates of TEs with abnormal secondary cell walls in total TEs that differentiated from cells electroporated with lambda dsRNA, Z2657 dsRNA or without dsRNA. Lambda dsRNA corresponding to a partial sequence of lambda DNA was used as a control.
(g) Rates of typical TE abnormalities in cultures treated with Z2657 dsRNA. Each data point represents the mean of three independent cultures ± SD.2

Pharmacological modification of cellulose synthesis of the secondary cell wall has previously been shown to cause dispersed lignification and less regularly shaped walls in Zinnia TEs (Taylor et al., 1992). Phloroglucinol staining of Z2657 dsRNA-treated TEs revealed an altered lignification pattern (weak and partial staining on secondary walls, and a subset of cells that showed ectopic lignification) compared with the fully stained normal TEs (Figure 5a–d). Normally stained TEs after the introduction of Z2657 dsRNA were reduced significantly compared with that of lambda dsRNA (Figure 5e).

Figure 5.

 Effect of Z2657 double-stranded RNA (dsRNA) on lignin staining of tracheary elements (TEs).
(a–d) Light micrographs of TEs with normal (a), weak (b), partial (c) and ectopic (d) lignin stainings. Scale bars = 25 μm.
(e) Rates of TEs with nomal lignin staining in total TEs that differentiated from cells electroporated with indicated dsRNAs. Lambda dsRNA corresponding to a partial sequence of lambda DNA was used as a control. Each data point represents the mean of three independent cultures ± SD.

To ensure that the observed phenotype resulted from the knocking down of the expression of a secondary cell wall-type CesA gene by Z2657 dsRNA-mediated silencing, we next carried out RNA silencing experiments targeted to two different types of Zinnia CesA genes by using Z632 and Z811 dsRNAs (Figures S2 and S3). Z632 is homologous to AtCesA2, AtCesA5, AtCesA6 and AtCesA9, of which AtCesA6 is thought to participate in primary wall formation (Desprez et al., 2002; Fagard et al., 2000). Z811 is homologous to AtCesA7, which is involved in secondary wall formation similarly to AtCesA8 (Taylor et al., 1999, 2000). Microarray analysis indicated that Z632 expression remains constitutively low throughout TE differentiation, whereas Z811 transcripts are specifically expressed at the onset of the secondary cell-wall formation (Figure S4; Demura et al., 2002). In addition, in situ hybridization revealed a differentiating TE-specific signal of Z811 in Zinnia plants (Demura et al., 2002). Introduction of Z632 dsRNA into Zinnia cells resulted in a slight increase in abnormal TEs compared with control lambda dsRNA-treated TEs, but was much less than that observed for Z2657 dsRNA cells (Figure 6a). In contrast, Z811 dsRNA showed a similar effect on TE abnormality as Z2657 dsRNA (Figure 6b,c). Although similar phenotypes were observed when introducing Z2657 and Z811 dsRNAs, a significant and specific reduction of the ZeCesA1 mRNA level (∼40%) was detected only after the introduction of the corresponding Z2657 dsRNA (Figure 7). The expression of ZCP4, a TE-specific cysteine protease (Figure S4; Demura et al., 2002; Pesquet et al., 2004) that has been suggested to play a role in TE cell death (Fukuda, 2000), was used as a reference to reflect TE gene expression. Together, these results indicate that modulation of Zinnia secondary cell-wall CesAs by dsRNAs caused TE abnormality, as expected from the phenotype observed in vessels of Arabidopsis whole plants with mutation in the equivalent CesA genes.

Figure 6.

 Comparison of the effects of double-stranded RNAs (dsRNAs) corresponding to different CesA genes of Zinnia on secondary cell-wall formation in tracheary element (TE) differentiation.
(a, b) Effect of Z632 dsRNA (a) and Z811 dsRNA (b) on secondary cell-wall formation of TEs was compared with that of Z2657 dsRNA.
(c) Rates of TE abnormalities in cultures treated with indicated dsRNAs. Lambda dsRNA corresponding to a partial sequence of lambda DNA was used as a control. Each data point represents the mean of three independent cultures ± SD.

Figure 7.

 Comparison of the effects of double-stranded RNAs (dsRNAs) on the accumulation of ZeCesA1 mRNA in differentiating tracheary elements (TEs) by quantitative RT-PCR analysis.
(a) Amplified products of ZeCesA1 (upper) and ZCP4 (bottom) after 28 cycles.
(b) Quantified expression levels of ZCP4 (white column) and ZeCesA1 (grey column). Each template cDNA was reverse-transcribed from RNA mixtures of three independent 80-h-old cultures electroporated with the indicated dsRNAs. Lambda dsRNA corresponding to a partial sequence of lambda DNA was used as a control. Each data point represents the mean of three independent PCRs ± SD. The experiment was repeated, yielding similar results.


One of the main difficulties with the transformation of single-celled plant systems is that the presence of a cell wall greatly reduces transformation efficiency. Traditionally, the solution to this problem has been to use protoplasts (Fromm et al., 1987). However, the preparation and use of protoplasts is at best problematic in Zinnia cell culture, because the time course of TE differentiation from Zinnia protoplasts is variable, and is usually delayed significantly, and the TE differentiation efficiency is very low compared with mesophyll cells with cell walls (Kohlenbach and Schöpke, 1981). To establish valuable transformation systems, it is important to keep TE differentiation as close to the native Zinnia system as possible, so that previously obtained data on morphological hallmarks and gene expression can be used.

Electroporation of intact plant cells has been successfully performed without preparing protoplasts (Koscianska and Wypijewski, 2001; Lin et al., 1997; Sabri et al., 1996; Wu and Feng, 1999). However, freshly isolated Zinnia mesophyll cells were recalcitrant to electroporation transformation. Because partial digestion of the cell wall greatly improved transformation efficiency, it appears that the Zinnia cell wall is an efficient barrier to electroporation-induced cell transformation. Pectin-degrading enzymes are generally used to increase access to cellular content without altering the visible cell-wall structure for in situ RT-PCR (Urbanczyk-Wochniak et al., 2002). In our case, it seemed that pectolyase mildly hydrolyzed primary cell-wall pectins, whereas cellulase and macerozymes probably released ‘partial protoplasts’.

The originality of our protocol is that it combines effective cell transformation with efficient TE differentiation. The optimal electrical parameters, we determined were in agreement with the general finding that low voltage and high capacitance are better for transformation of plant cells (Table 2) (Fromm et al., 1987). Several other variable parameters, such as the use of KCl rather than NaCl, and the addition of HEPES, CaCl2, PEG and salmon carrier DNA, also improved the efficiency of our system.

In order to broaden the utility of the Zinnia system, we adapted the transformation protocol for use with a number of common cellular and molecular biological techniques. The first of these was protein subcellular localization. We were able to target fluorescent proteins in the cytoplasm and the nucleus successfully (Figure 2). One potential use of this system is to tag subcellular compartments fluorescently, and thereby allow direct monitoring of the modification of organelle morphology during TE differentiation. Up to now, these observations have been performed using dyes, which are less specific and can have toxic effects on the cell culture. This technique could also be extended to the study of protein trafficking during TE differentiation, allowing protein maturation and subcellular localization to be monitored over the time course of TE differentiation.

The protocol was further extended by co-expressing two distinct fluorescent proteins derived from the co-transformation of two different plasmids (Figure 2). This has potential in the field of protein–protein interaction studies within the developmental context of TE formation. Using techniques such as Fluorescence Resonance Energy Transfer imaging and/or Bimolecular Fluorescence Complementation or ‘split YFP’ (Bhat et al., 2006), protein–protein interactions could be monitored in differentiating Zinnia cells, providing accessible biochemical evidence of the interactions in a physiologically relevant setting.

We were also able to monitor promoter activity directly during TE differentiation. PTED3::YFPnuc expression observed during TE differentiation was in good agreement with expression patterns of TED3 transcripts previously described (Demura and Fukuda, 1994; Figure S4). It is interesting to note that an absence of significant PTED3::YFPnuc expression in differentiating TEs might suggest TED3 transcription was tightly restricted to cells in a TE precursor-like state in the Zinnia system. This transformation method also provides the possibility for efficient promoter deletion studies in a physiologically relevant system.

Finally, and most importantly, we have developed the transformation protocol for dsRNA-mediated silencing assays. As dsRNAs were simply transcribed in vitro from cDNA clones that had been already constructed for microarray analysis, this provides an efficient way to produce a large number of different dsRNAs, enabling one to perform RNA silencing analysis systemically and comprehensively without labour-intensive constructions. In addition, plasmids can be used in combination with dsRNAs to monitor the effect of gene silencing (with dsRNA) on reporter protein expression and/or localization (with plasmid DNA) at the single-cell level.

Genes encoding the catalytic subunit of cellulose synthase, CesA, were efficiently silenced by introduction of CesA dsRNAs. CesAs were chosen to estimate selective suppression by dsRNA-mediated silencing because they are involved in cell-wall biosynthesis in Arabidopsis, and we expected that any perturbation of CesA function would be easily scorable in a phenotypic screen in the Zinnia system. Our assays confirmed the functionality of the system and the functional conservation of CesA genes between plant species. Both Z2657 and Z811 dsRNAs affected secondary wall formation by either reducing its light refringency/apparent cellulose content (leading to ‘thin’ TEs), and/or affecting the link between primary and secondary walls (leading to ‘separated’ TEs). Based on sequence similarity to known CesA genes (Figures S2 and S3), and on specific expression during the onset of secondary cell-wall formation (Figure S4), they were both expected to have a similar function to their putative Arabidopsis orthologues, AtCesA8 and AtCesA7, which have been shown to be involved in secondary cell-wall biosynthesis. The results presented here suggest that Z2657 and Z811 can be considered as ‘true’ functional orthologues of AtCesA8 and AtCesA7, respectively.

Although we still do not know how many cells are affected by dsRNA-induced silencing, the fact that Z2657 and Z811 dsRNAs increased abnormal TEs suggests that the quantity of dsRNA used in this study was sufficient to ensure RNA silencing throughout the differentiation process. Because RNA silencing results in various levels of inhibition of the target gene expression in different individuals (for example, for CDC45 modulation in Arabidopsis; Stevens et al., 2003), the observed 40% overall reduction in RNA accumulation would most likely result from various degrees of inhibition of ZeCesA1 gene expression in individual cells (Figure 7). The extensive downregulation of our target gene expression shows that direct dsRNA introduction is more efficient than reporter plasmid transformation. This can be explained by both the smaller size of the dsRNAs allowing cell penetration and that they have a cytoplasmic site of action, whereas reporter plasmids would need to be transported to the nucleus to be transcribed.

It has been suggested that secondary cell-wall CesA proteins are necessary during TE differentiation, to form thick and tight secondary walls that guide lignin deposition. Our data further confirms the guidance role of cellulosic secondary cell walls on lignification, because reduced and ectopic lignification patterns were specifically observed in Z2657 dsRNA-treated cells (Figure 5). In Arabidopsis irx mutants, the inward collapse of abnormal TEs is caused by negative pressure generated by water transport in plants (Turner and Somerville, 1997). We did not observe any collapse in our abnormal TEs in either the Z2657 or Z811 dsRNA cultures, reinforcing the explanation for the Arabidopsis phenotype. Finally, as we have observed similar phenotypes in TEs of both Arabidopsis irx mutants and Z2657 and Z811 dsRNA-treated Zinnia cells, we can conclude that the irx phenotype in whole Arabidopsis plants is a primary effect, rather than a secondary effect of the irx mutation.

The application of our transformation protocol is expected to greatly increase the utility of the Zinnia system and the study of TE formation. As outlined here, the Zinnia system is a stand-alone system of great value not only for the study of TE formation, but also for the study of secondary cell-wall formation. It is also of considerable use in verifying results observed in whole plant systems. On the other hand, it can also be of use in cases where different phenotypes are observed in model plants and in Zinnia TEs. Such differences may provide clues to the multiple roles of genes during differentiation processes. Moreover, this system is potentially of great value in cases where loss-of-function is lethal. As one of the major challenges in all systems is to gain an understanding of the biological function of unknown genes, our method is expected to be of considerable use, particularly in the study of TE differentiation.

Experimental procedures

Plant material and culture conditions

Seedlings of Zinnia (Z. elegans L. cv. Canary bird; Takii shubyo, were grown on vermiculite for 2 weeks to harvest first leaves. Mesophyll cells were isolated in FK medium, containing 0.1 mg l−1 of naphthylacetic acid and 0.2 mg l−1 of benzyladenine, according to the method described by Fukuda and Komamine (1980). After electroporation, cells were cultured under hypertonic culture conditions: FK medium with 50 mm higher content (finally 250 mm) in mannitol.

Preparation of dsRNAs

For the preparation of GFP dsRNA and P35S dsRNA, DNA fragments from +1 to +400 and from −467 to −68, relative to the site of initiation of translation, were amplified from pGFP2(GA)5II (Dong et al., 2001; Kost et al., 1998), respectively, and subcloned into pGEM-T Easy vector (Promega, A 0.4-kbp fragment from the middle of the GUS open reading frame (ORF) was subcloned into pGEM-T Easy vector. A PvuII fragment of 532 bp of lambda DNA (Takara Bio, was subcloned into the pGEM-5Zf(+) vector (Promega) at the EcoRV site, for preparation of lambda dsRNA. The Zinnia clones (Z2657, Z632 and Z811) have already been subcloned into pGEM-T Easy vector in the construction of the equalized cDNA library (Demura et al., 2002). Templates for in vitro transcription were amplified from each vector by using KOD plus (Toyobo, with M13 forward and reverse primers. AmpliScribe T7 and SP6 high-yield transcription kits (Epicentre, were used to produce both RNA strands from the each template, according to the manufacturer’s instructions. After transcription, both RNA strands were mixed to 4 mg ml−1 in H2O and denatured at 95°C, followed by 65°C for 10 min and then 42°C for 10 min. Annealed RNAs were confirmed on a non-denaturing electrophoresis and then compared with non-annealed RNAs.

Introduction of plasmid DNA/dsRNA into Zinnia cells by electroporation

Pre-treatment of freshly isolated Zinnia mesophyll cells and electroporation were performed using the following solutions: a pectolyase solution [0.36 mg ml−1 pectolyase Y-23 (Kikkoman,, 11 mm 2-N-morpholinoethanesulfonic acid, pH 5.5, 330 mm mannitol and 18 mm CaCl2], a washing solution (260 mm mannitol, 5 mm CaCl2 and 5 mm HEPES, pH 7.2) and a KCl/PEG solution (2 m KCl:40% PEG 6000 = 1 : 10). Zinnia mesophyll cells were centrifuged to above 3 × 106 cells ml−1 in the normal culture medium, and were then mixed gently with a 1/2.7 volume of the pectolyase solution. After incubation at 30°C for 30 min, the cells were washed twice with the washing solution and resuspended in the washing solution at 3–5 × 106 cells ml−1. Then, 700 μl of the cell suspension was mixed with 40 μl of 2 mg ml−1 plasmid DNA and/or 5 μl of 4 mg ml−1 dsRNA followed by 110 μl of the KCl/PEG solution in a 0.4-cm-gapped electroporation cuvette. Electroporation was then performed using a Gene Pulser II with a Capacitance Extender Plus (Bio-Rad, at 325 V cm−1 and 900 μF, yielding the time constant of 45–60 msec at room temperature (20–25°C) under optimal conditions. After electropulsation, the cells were immediately transferred into the hypertonic culture medium at about 11 × 104 cells ml−1.


Cells electroporated with P35S::GUS (p35SGUS; Ito and Fukuda, 2002) and then cultured were fixed in the hypertonic culture medium supplemented with 0.2% glutaraldehyde and X-Gluc at 4°C, and were then incubated in an X-Gluc solution (1 mm X-Gluc, 50 mm sodium phosphate, pH 7.0 and 20% methanol) at 37°C for 24 h. For detection of lignin (Pomar et al., 2002), cell suspensions were incubated with an equal volume of 1% phloroglucinol dissolved in 18% HCl at room temperature.

Fluorometric assay

Cells cultured for 48 h were ultrasonically homogenized in an extraction solution (50 mm sodium phosphate, pH 7.0, 10 mm EDTA, 10 mm 2-mercaptoethanol and 20 mg ml−1 polyvinylpolypyrrolidone). GUS activity in the extract was measured using 4-methyl-β-umbelliferyl glucuronide according to the method described by Jefferson et al. (1987). Dilutions of a solution of methylumbelliferone were used as standards.


TE differentiation (%) was measured as the percentage of TEs per total living cells and TEs. Zinnia cells were stained with 50 μm Evan’s blue just before observation, to determine cell viability. To detect cellulose in secondary cell walls of developing TEs, cells were stained using 0.01% calcofluor as described previously (Pesquet et al., 2005). At least 200 TEs were observed in each culture for the determination of rates of abnormal TEs and normally stained TEs by phloroglucinol. GFP- and YFP-expressing cells (%) were measured as the percentage of cells showing specific fluorescence per total living cells and TEs. CFP/YFP co-expressing cells (%) were measured as the percentage of cells showing CFP and YFP expression at the same time. The ZeTED3 promoter fragment (Igarashi et al., 1998) was subcloned into pBGYN (Kubo et al., 2005) and used for nuclear targeting analysis and promoter activity analysis. For the co-expression experiment, pY35GS and pC35GS plasmids (M. Kubo and T. Demura, RIKEN, Yokohama, Japan; unpublished data) were used, which harboured P35S::YFP-P35S::GATEWAY (Invitrogen, and P35S::CFP-P35S::GATEWAY, respectively, on the pBluescript II cloning vector (Stratagene,

RT-PCR analysis

Cells were collected by centrifugation, pulverized in liquid nitrogen, dissolved in equal parts of an extraction buffer (200 mm Tris–HCl, pH 9.0, 100 mm NaCl, 10 mm EDTA, 0.5% SDS and 14 mm 2-mercaptoethanol) and phenol saturated with 10 mm Tris–HCl, 1 mm EDTA, pH 8.0. Chloroform:isoamyl alcohol (24:1) was then added, mixed and centrifuged. Aqueous supernatant was collected and mixed once more with chloroform:isoamyl alcohol. RNA was precipitated by LiCl (final concentration, 2 m) and resuspended in a Tris–EDTA solution (10 mm Tris–HCl and 1 mm EDTA, pH 8.0). Removal of polysaccharides was performed by the addition of 300 mm sodium acetate, followed by centrifugation. RNA of the supernatant was precipitated by the addition of 2.5 volumes of ethanol, resuspended in the Tris–EDTA solution and treated with RNase-free DNaseI (Roche, cDNA synthesis was performed with oligo (dT) using SuperScript II RNase reverse transcriptase (Invitrogen). PCR was performed with LightCycler-Faststart DNA Master SYBR Green I (Roche). The reaction included touchdown cycles from 61°C to 55°C in the presence of 4.0 and 4.5 mm MgCl2 for ZCP4 and ZeCesA1, respectively. The primers used were: 5′-GTGGATTTGCCAAAATCGGTTGATT-3′ and 5′-ATTGCACCCGTTGTTGAAGGTTGT-3′ for ZCP4, and 5′-TATTGGGTTTGGTCCTGATGGGAA-3′ and 5′-GTTACAGGCTCCTTTTCCATCTCA-3′ for ZeCesA1.


We thank Dr Brian Jones for his critical reading of this manuscript and Dr Nam-Hai Chua for providing pGFP2(GA)5II. This work was supported in part by Grants-in-Aid from the Ministry of Education, Science, Sports and Culture of Japan (14036205), and the Japan Society for the Promotion of Science (15770043, 17207004).