ISOLATION OF PROTOPLASTS AND VACUOLES FROM … · Protoplast isolation from stem parenchyma cells The green peripheral part of the sugarcane stem (or culm), where vascular bundles

Gnanasambandam, A. & Birch, R.G. Proc. Aust. Soc. Sugar Cane Technol., Vol. 28: 2006

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ISOLATION OF PROTOPLASTS AND VACUOLES FROM SUGARCANE SUSPENSION AND STEM PARENCHYMA CELLS

By

ANNATHURAI GNANASAMBANDAM and ROBERT G. BIRCH

Department of Botany, The University of Queensland, St. Lucia, Qld Contact author: [email protected]

(Present address: BSES Limited, Indooroopilly, Australia)

KEYWORDS: Protoplasts, Vacuoles, Metabolic Engineering.

Abstract THE VACUOLE is a compartment within sugarcane cells in which most of the sucrose is stored. Despite its importance, the sugarcane vacuole is not well understood, because it has been very difficult to examine from sucrose-accumulating stem cells. It is only separated from other cell components by a single membrane which is generally broken during attempts to isolate the vacuole for study. One new approach to sugarcane improvement is to direct novel enzymes into the vacuole to convert sucrose into higher value biomaterials. Unfortunately, the vacuole is very acidic and it also contains proteolytic enzymes that seem to degrade most introduced enzymes before they can perform useful bioconversions. This new potential led us to revisit the challenge of vacuole isolation from mature sugarcane stems. Our immediate interest was to use the isolated vacuoles to understand how to stabilise introduced enzymes. They would also be very useful in studies to understand how sugarcane achieves its exceptional stored sugar concentrations. A first step to vacuole isolation is the removal of the cellulose walls surrounding sugarcane cells, yielding protoplasts. So we developed a method to isolate protoplasts from sugarcane immature and mature stem parenchyma cells by: (i) using an enzyme solution containing cellulase onozuka RS, macerozyme, pectinase and driselase; (ii) adjusting the solution to match the internode osmotic strength; (iii) pretreating internodes at 40C for 1�2 days before the enzyme treatment; (iv) vacuum infiltration of tissues in enzyme solution and changing the solution after 1�2 hours of incubation; (v) purification of protoplasts in Ficoll gradients. However, osmotic lysis of these protoplasts yielded less than 2% of vacuoles in most trials and polybase-induced lysis did not yield any vacuoles.

Introduction

Sugarcane accumulates sucrose at concentrations up to 62% dry weight (16�25% fresh weight) in the storage parenchyma cells of the mature culm. A large vacuole found in these storage cells contains most of the stored sucrose (Moore et al., 1997). Hence, the ability to direct foreign proteins to the sugarcane vacuole and retain biological activity of such proteins in the vacuole is of substantial practical relevance to the sugarcane industry to synthesise valuable novel products and compounds other than sucrose.

There is no published method for isolation of vacuoles from sugarcane mature stem parenchyma cells, and their physiology is not well understood. For example, the nature of any proteolytic enzymes (enzymes that may inactivate introduced proteins) in the sucrose storage


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vacuoles has not been reported. Hence, we would like to isolate vacuoles from sugarcane stem parenchyma cells for characterisation of proteolytic enzymes and for compartmentation analysis to identify and characterise foreign proteins that are targeted to the vacuole.

The first step towards vacuole isolation is generally to isolate intact cells from which the cellulose walls have been removed. These wall-free cells, called protoplasts, are generally obtained by treatment of plant tissues with enzymes to dissolve the cell walls.

Methods to isolate protoplasts from sugarcane cells grown in tissue culture are well established (Srinivasan and Vasil, 1986; Taylor et al., 1992; Matsuoka et al., 1996; Aftab and Iqbal, 1999). However, only a few reports are available for isolation of protoplasts from sugarcane plants; and these are limited to young tissues such as the spindle region (Krishnamurthy, 1973) and immature stalk tissue (Thom et al., 1983; Maretzki and Thom, 1987). Unfortunately, mature tissues generally develop hardened cell walls which are more difficult to remove without damaging the cells.

Here we report a method for efficient isolation of protoplasts from sugarcane mature stem parenchyma cells, as a step towards the isolation of the vacuoles from these cells. Materials and methods

Embryogenic callus and cell suspension culture Embryogenic callus and cell suspension cultures of commercial sugarcane cultivar

Q63 were initiated and maintained as described by Franks and Birch (1991) and Taylor et al. (1992), respectively.

Solutions used 1. Enzyme solution A: 2% cellulase onozuka RS, 0.2% pectinase, 0.1%

driselase and 0.2% macerozyme in wash solution A [pH 5.9, 3 mM MES and 0.7 mM NaH2PO4 H2O with 0.2 M each of mannitol and sorbitol].

2. Enzyme solution B: 2% cellulase onozuka RS, 0.2% pectinase, 0.1% driselase and 0.2% macerozyme in wash solution B (wash solution A with 0.25 M each of mannitol and sorbitol).

3. Enzyme solution C: 2.5% cellulase onozuka RS, 0.2% pectinase, 0.1% driselase and 1% macerozyme in wash solution C (wash solution A with 0.3 M each of mannitol and sorbitol).

4. Lysis buffer A or B or C: 0.1 or 0.2 or 0.13 M KH2PO4/K2HPO4, respectively, 2 mM EDTA-Tris (pH 8) and 2.5 mM DTT.

5. Lysis buffer D: 0.2 M sorbitol, 10% Ficoll 400 and 10 mM Hepes-KOH (pH 7.5).

Protoplast enzyme solution The enzyme solution used for isolation of protoplasts was prepared as described by

Maas et al. (1995) and Dombrowski et al. (1994). The enzymes [cellulase onozuka RS (from mutant strain of Trichoderma viride) and macerozyme (from Rhizopus sp.) (Yakult Manufacturing Co., Japan); pectinase (from Aspergillus niger) and driselase (from Basidiomycetes sp.) (Sigma)] were slowly dissolved in millipore-filtered sterile water and stirred for 2 to 3 hours at 240C.

Care was taken not to create foaming, which may result in the inactivation of the enzymes in solution. Then, CaCl2, NaH2PO4, MES, mannitol and sorbitol were added in that order as required from stock solutions, and the pH was adjusted to 5.9.

When dissolved, the enzyme solution was slowly filter sterilised using a 0.22 !m filter, and stored in 20 mL aliquots at �200C.


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Protoplast isolation from suspension cells Protoplasts were isolated from suspension culture cells as described by Srinivasan

and Vasil (1986). Five mL of the suspension was mixed with 25 mL of filter sterilised enzyme solution A in a Petri dish and incubated on a shaker for 4 to 5 hours at 60 r/min in darkness at 270C.

Protoplasts were filtered through two 200 !m nylon mesh [Swiss Screens (Australia) Pty. Ltd., Brisbane, Australia], pelleted by centrifugation at 100 g for 5 min and washed three times in wash solution A.

Protoplast isolation from stem parenchyma cells The green peripheral part of the sugarcane stem (or culm), where vascular bundles are closely packed, was peeled off. Internode cores were then cut into 1�2 mm thick slices in Petri dishes containing the wash solution B or C.

Tissue pieces were then transferred to the enzyme solution B or C and incubated for 1 to 4 hours on an orbital shaker at 60 r/min in darkness at 270C. Internode numbering was done as described in Moore (1987) such that numbers increase down the stem, with the internode below the node attached to the top visible dewlap leaf being designated as 1.

For internode one, enzyme solution B and wash solution B were used. For internodes 2 to 18, enzyme solution C and wash solution C were used. For storage at 40C, stem internodes were kept for 24 hours or more in a polythene bag.

After tissue digestion, released protoplasts were filtered through two 200 !m nylon mesh over two layers of cheese cloth, washed in wash solution B or C by sedimenting at 100 g for 5 min. The protoplasts were further purified twice on Ficoll 400 (Pharmacia) gradients using the procedure of Boller and Kende (1979) as given in Wagner (1987).

Protoplasts were gently mixed with one volume of 15�20% (w/v) Ficoll prepared in wash solution B or C. This solution is overlayered with 8, 6 and 0% (w/v) Ficoll zones (10 mL), each containing mannitol and sorbitol and buffers as in wash solution B or C.

After centrifugation at 1000 g for 10�20 min at 40C, purified protoplasts were gently collected from the 0�6% interface and diluted into wash solution B or C and collected from this suspension by sedimentation at 100 g for 5 min.

Isolation of vacuoles Osmotic lysis of protoplasts involved rapid (5 sec) dilution of 1 or 2 mL suspension

of protoplasts in wash solution A or B or C in a round-bottomed glass tube with 10 or 20 mL of lysis buffer A/B/C/D. The suspension was gently stirred (about 2 min) or pipetted up and down using a glass pipette to effect emergence of vacuoles from lysing protoplasts (Wagner, 1987).

In polybase-induced lysis of protoplasts, washed protoplasts were layered onto a three-step gradient containing the following: first step, 5 mL medium of 25 mM Tris-MES buffer, pH 6.5, 0.6 M mannitol containing Ficoll 2%, DEAE dextran 4 mg/ml; second step, 2 mL medium of 25 mM Tris-MES buffer, pH 8.0, 0.6 M mannitol (medium D) containing Ficoll 5%, dextran sulfate 4 mg/ml; third step, 2 mL medium D containing Ficoll 20%.

The gradient was centrifuged for 30 min at 2000 g at 170C and the unlysed protoplasts were collected at the interface between the second and third steps (Wagner, 1987).

Neutral red staining Fifteen !L of 10 mg/mL neutral red was added per 12 mL of wash solution

containing the protoplasts to visualise the vacuoles. Sugarcane suspension cells and friable


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calli cells (obtained from suspension cells) were suspended either in MSC3 medium or 50 mM sodium phosphate buffer (pH 7) for neutral red staining. Results

Examination of sugarcane suspension cells, including staining to identify vacuoles We attempted to visually distinguish cytosolic and vacuolar regions in suspension

cells. When sugarcane Q63 cells were examined without any staining, the distinction between the vacuole and cytosol was visible only in a few cells (Figure 1A).

In a few cells, the nucleus was visible (arrow in Figure 1C). Hence, to confirm the vacuolar region, neutral red staining was used.

When the Q63 suspension cells in MSC3 medium were stained with the neutral red, the dye did not stain the vacuoles, probably due to low pH of the medium. In some of the cells, only the nucleus (arrow in Figure 1D) and the nucleolus (dotted arrow in Figure 1D) were preferentially stained.

However, when the cells were stained after transferring them to 50 mM sodium phosphate buffer (pH 7.0), the vacuoles accumulated the neutral red readily and the cytosolic and the vacuolar regions were clearly identified in many cells (Figure 1B). In these cells, the vacuoles occupied more than 80% of the cell region.

Isolation of protoplasts from suspension cells Protoplasts were successfully isolated from sugarcane suspension cells using enzyme

solution A (Figure 1E). In these protoplasts, vacuoles were visible without any staining under phase contrast microscope (Figure 1E).

With neutral red staining, most of the vacuoles accumulated the dye and appeared pink to red (Figure 1F). However, some protoplasts showed a reversed staining pattern, with pink to red cytosol and unstained or faintly stained vacuole (dotted arrows in Figure 1F).

Isolation of vacuoles from suspension cell protoplasts Isolation of the vacuoles from suspension cell protoplasts using osmotic and

polybase-induced lysis proved to be difficult. Several attempts using polybase-induced lysis could not yield any vacuoles. In contrast, osmotic lysis of protoplasts yielded a few vacuoles in most of the trials (arrows in Figure 1F and I).

However, in this method, addition of 10 mL of lysis buffer A to the protoplasts could not lyse most of the protoplasts (Figure 1F), while addition of 20 mL of lysis buffer A caused bursting of the majority of the protoplasts (Figure 1I). Reproducible and efficient lysis of most of the protoplasts to yield vacuoles was not accomplished.

Isolation of protoplasts from stem cells When slices from immature internode cores of sugarcane cultivars Q63, Q117, Q155

and Pindar were incubated in enzyme solution A (used for isolating protoplasts from cell suspension culture), a lot of cell debris was found after 4 hours.

The pH of the solution (pH 5.9) in the three replicate plates had dropped to 2.77, 3.86 and 3.69 indicating lysis of cells. Increasing the mannitol and sorbitol concentration from 0.2 M to 0.25 M in the enzyme solution (enzyme solution B) yielded protoplasts from internode 1 for all four cultivars.

Within 2 hours of incubation in enzyme solution B, all the tissues were completely digested and yielded the protoplasts (Figure 1H, K and L).

In these protoplasts, neutral red staining indicated that the vacuole has occupied more than 90% of the cell volume (Figure 1K). In some protoplasts, the nucleus can be identified (arrow in Figure 1L).


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Fig. 1�Protoplasts and vacuoles from sugarcane suspension and stem parenchyma cells. Sugarcane suspension cells without any staining (A and C) and with neutral red staining (B and D). Nuclei are shown by black arrows in C, D and L and nucleolus is shown by dotted arrow in D. Protoplasts from suspension cells without any staining (E � phase contrast micrograph) and with neutral red staining (F and I); Isolated vacuoles are shown by black arrows in F and I. Protoplasts showing a reverse staining pattern, with stained cytosol and unstained or faintly stained vacuole are shown by dotted arrows in F. Protoplasts (H, K-L) and vacuole (M) from sugarcane stem cells (G � unstained transverse section). Protoplasts after polybase induced lysis (J). Bar = 25 mm (except in B and K-M where it equals 5 mm).


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Efficient isolation and purification of protoplasts from internodes 2 and 3 of the four cultivars was obtained by:

(i) Increasing the mannitol and sorbitol concentration to 0.3 M each and increasing the cellulase and macerozyme concentrations to 2.5% and 1%, respectively (enzyme solution C).

(ii) Slicing the tissues under wash solution C and vacuum infiltration of the tissues with enzyme solution C at the start of the incubation (Upon slicing, the spaces between the cells are filled with air that hinders the digestive enzymes coming into contact with the cell walls. This difficulty is avoided by infiltrating the tissues under vacuum or by slicing the tissue under wash solution, draining the strips and transferring them to the incubation medium).

(iii) Replacing old enzymes with new enzyme solution after 1 or 2 hours of incubation.

(iv) Purification of protoplasts in Ficoll 400 gradients. Incubation of stem tissues (Figure 1G) from internode 4 in enzyme solution C

yielded no protoplasts. However, storage of stem internodes in a polythene bag at 40C for 24 hours or more prior to enzyme digestion, yielded protoplasts with enzyme solution C. The stem segments stored in this way were able to yield protoplasts for up to 3 weeks in enzyme solution B (for internodes 1 to 3) or C (for internodes 4 to 18).

Attempts to isolate vacuoles from stem protoplasts Some of the trials attempted for vacuole isolation from stem cells of cultivars Q63,

Q117, Q155 and Pindar are given here. A flotation technique was used for polybase-induced lysis of protoplasts from stem cells as the protoplasts required flotation during purification in Ficoll gradients. In contrast, a sedimentation technique was used for protoplasts from suspension cells.

Trial 1: The protoplasts were chilled on ice for 30 minutes and were subjected to osmotic lysis using lysis buffer C. Addition of 10 mL of lysis buffer C at 270C or 380C yielded less than 1% of vacuoles/protoplasts. Addition of 10 mL of lysis buffer C at 550C did not yield any vacuoles.

Trial 2: Addition of 10 mL or 20 mL of lysis buffer A resulted in less than 1% of vacuoles/protoplasts. The yield of vacuoles increased to 2% of vacuoles/protoplasts when 20 mL of prewarmed lysis buffer A at 420C was added. No vacuoles were obtained when 20 mL of lysis buffer B was added.

Trial 3: Polybase-induced lysis of protoplasts was tried several times with no success of isolating vacuoles (Figure 1J).

Trial 4: The method of Dombrowski et al. (1994), involving addition of lysis buffer D, yielded less than 1% of vacuoles/protoplasts (Figure 1M). Discussion

Isolation of protoplasts from stem parenchyma cells The procedure described here allows isolation of protoplasts from stem parenchyma

cells of sugarcane. Conditions optimised using one cultivar, Q117, were readily adapted to three more cultivars, Q63, Q155 and Pindar, indicating that the procedure can probably be extended rapidly to additional sugarcane cultivars.

Though protoplasts of many crops are generally isolated at 25�300C, sensitive and difficult tissues benefit from long-term isolations at low temperatures (7�120C). Prolonged


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incubation of cells in enzyme solution at low temperature (140C) followed by a short time at about 300C have proved useful in isolation of protoplasts from cereal species (Potrykus et al., 1977; Vasil and Vasil, 1980).

To isolate protoplasts from sugarcane suspension cells, Srinivasan and Vasil (1986) used an incubation period of either 5 hours at 270C or 16 hours at 150C. However, prolonged incubation in enzyme solution may affect the quality of protoplasts and isolation of vacuoles. Hence, a pre-treatment at 40C was tried for the stem segments and was found to be required for isolation of protoplasts from internodes 4 to 18 of sugarcane.

This cold treatment might have reduced the osmotic potential of the mature cells, enabling them to yield protoplasts in 0.3 M sorbitol and 0.3 M mannitol. It might have also changed the cell wall composition to facilitate the attack of enzymes used.

Growing plants or cells in the dark for a few days before isolation of protoplasts seems to reduce osmotic pressure, and is reportedly helpful in the isolation and culture of protoplasts (Vasil and Vasil, 1980).

Storage of stem internodal segments in a polythene bag at 40C enabled the isolation of protoplasts for two to three weeks. This is helpful, as it guarantees a continuous supply of protoplasts for an extended period of time from a single plant.

This would be particularly useful for preliminary experiments, for example, for optimising vacuole isolation or enzyme analysis. However, the cold treatment may be a limitation, as it could cause stress and change the physiology of the stem cells.

The conditions determined in this study provide an ideal starting point for establishing a procedure for isolating protoplasts without a cold treatment. To develop a procedure without a cold treatment, higher osmotic concentrations of sorbitol and mannitol should be tried in the first instance.

In this study, the sorbitol/mannitol was increased from 0.5 M for internode 1 to 0.6 M for internodes 2 to 18 to get the protoplasts, indicating that higher osmotic concentrations above 0.6 M mannitol/sorbitol could be helpful to avoid the cold treatment.

Once the osmotic concentrations are optimised, higher concentrations of enzymes or more powerful enzymes (like pectolyase Y23) could be used. Cellulase onozuka RS used in this study is a more powerful enzyme than cellulase onozuka R10 enzyme used by Krishnamurthy (1973) and Maretzki and Thom (1987) to isolate protoplasts from immature stem cells. It is capable of dissolving walls of a wider range of plant tissues including those of the bundle sheath of several C4 plants. Among the pectinases, pectolyase Y23 appears to be most useful, even with otherwise intractable tissues (Wagner, 1987).

In sugarcane, pectolyase Y23 resulted in higher yields of protoplasts from both heterogeneous and homogeneous cell suspension cultures as compared to other pectinases (Taylor et al., 1992). Hence, cellulase onozuka RS with pectolyase Y23 and driselase could be tried with sugarcane stem parenchyma cells for refinement of the methods used in this study.

Isolation of vacuoles The choice of a vacuole isolation procedure depends on the nature of the plant

material or on the purpose of the scientist. (i) Osmotic lysis relies upon altering the osmotic conditions of the

protoplasts by rapid addition of a medium of lower osmotic concentration to the protoplast preparation. With a correct solute concentration difference between cells and added medium, breakage of the plasma membrane while maintaining an intact tonoplast (membrane enclosing


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vacuole) can be achieved. This is the most widely used method for vacuole isolation of many species.

(ii) Polybase-induced lysis relies on altering the charge on the external surface of the plasma membrane by the addition of DEAE dextran (which permeabilises the plasma membrane), followed by rapid removal of excess DEAE with dextran sulfate.

(iii) Lysis of protoplasts by shear under isotonic conditions involves selective rupture of plasma membrane when protoplasts are subjected to a high centrifugal force.

When vacuoles are released from protoplasts, irrespective of the procedure used, certain principles must be observed:

(i) Rapid protoplast isolation procedure to avoid prolonged exposure to wall-degrading enzymes.

(ii) Gentle lysis of the protoplasts in order to avoid rupture of the released vacuoles.

(iii) Efficient lysis in order to limit the occurrence of residual protoplasts which exhibit similar size and density to the vacuoles and are difficult to discard by centrifugation methods.

(iv) Rapid vacuole isolation, and limitation of the purification steps, in order to reduce bursting of the released vacuoles and leakage of solutes through the tonoplast (Boudet and Alibert, 1987).

In this study, polybase-induced lysis did not yield any vacuoles in any of the trials. The osmotic lysis method yielded less than 2% of vacuoles, based on the initial number of protoplasts in most of the trials.

This method works most successfully with protoplasts that are quite uniform in terms of osmotic potential, because lysis is produced by water-driven expansion of the protoplast, leading to stretching of the plasma membrane until it ruptures. At the lowered osmotic potentials required for maximum protoplast lysis, water also enters the vacuole, causing dramatic expansion and stretching and potentially also rupture of the tonoplast membrane (Leonard, 1987).

Thom et al. (1982) reported that the osmotic lysis method was unsatisfactory for sugarcane cells, because large numbers of protoplasts (30% or greater) contaminated the preparation, and resuspension of protoplasts in K2HPO4 (pH 8) increased the size of the tonoplast by about 100%.

The shear lysis method was therefore used to preserve the integrity of the tonoplast so that meaningful studies of transport could be made (Thom and Maretzki, 1985). However, Wagner (1987) reported that vacuoles prepared by this method were more highly contaminated (~10�30% of total extra-vacuolar marker activity) than those obtained using the osmotic or polybase-induced lysis.

Wagner (1987) suggested that the shear lysis approach is not ideal for compartmentation analysis where purity is very important, but it might be advantageous for transport studies where tonoplast integrity is important.

Thus, all the methods presently used to isolate vacuoles suffer from drawbacks of one kind or another. The yield of vacuoles is less than 40% with ideal starting tissue in most of the methods, and much lower with difficult tissues as in the present study.

The addition of chemicals such as EDTA or EGTA could make the plasma membrane highly susceptible to breakage but leaving the vacuolar membrane intact


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(Maretzki and Thom, 1989). Lysis of plasma membrane by digitonin to isolate vacuoles in plants (Le-Quoc et al., 1987) could be tried. With increasing knowledge on the constituents of plasma membrane and tonoplast, it could be possible in the future to lyse the plasma membrane by enzyme treatment without affecting the tonoplast. Conclusion

We have developed a method to isolate protoplasts from sugarcane immature and mature stem parenchyma cells, as a step towards isolation of the vacuoles from these cells. For mature stem cells, a pre-treatment of internodes at 40C for 1�2 days was required for protoplast isolation.

To avoid the cold stress and associated changes in the physiology of the stem cells, it may be possible to use more powerful protoplast releasing enzymes in a solution correctly adjusted to match the internode osmotic strength. Vacuole isolation was problematic, and osmotic lysis of protoplasts yielded less than 2% of vacuoles. Addition of chemicals such as EDTA, EGTA to weaken the plasma membrane or lysis of plasma membrane by digitonin could be tried to improve the vacuole yield in sugarcane. Acknowledgements

We thank the University of Queensland for Postgraduate Research Scholarships (OPRS and UQPRS) to AG.

REFERENCES Aftab, F. and Iqbal, J. (1999). Plant regeneration from protoplast derived from cell

suspension of adventive somatic embryos in sugarcane (Saccharum spp. hybrid cv. CoL-54 and cv. CP-43/33). Plant Cell Tissue Organ Cult., 56 (3): 155�162.

Boller, T. and Kende, H. (1979). Hydrolytic enzymes in the central vacuole of plant cells. Plant Physiol., 63: 1123�1132.

Boudet, A.M. and Alibert, G. (1987). Isolation of vacuoles and tonoplast from protoplasts. Methods Enzymol., 148: 74�87.

Dombrowski, J.E., Gomez, L., Chrispeels, M.J. and Raikhel, N.V. (1994). Targeting of proteins to the vacuole. In: Gelvin, S.B., Schilperoort, R.A. and Verma, D.P. S. ed.. Plant Molecular Biology Manual. Kluwer Academic Publishers, Belgium, J3: 1�29.

Franks, T. and Birch, R.G. (1991). Gene transfer into intact sugarcane cells using microprojectile bombardment. Aust. J. Plant Physiol., 18: 471�480.

Krishnamurthy, M. (1973). Isolation, fusion and multiplication of sugarcane protoplasts and comparison of sexual and parasexual hybridisation. Euphytica, 25 (1): 145�150.

Leonard, R.T. (1987). Preparation of tonoplast vesicles from isolated vacuoles. Methods Enzymol., 148: 82�89.

Le-Quoc, K., Le-Quoc, D. and Pugin, A. (1987). An efficient method for plant vacuole isolation using digitonin for plasmalemma lysis. J. Plant Physiol., 126: 329�335.

Maas, C., Reichel, C., Schell, J. and Steinbi", H.H. (1995). Preparation and transformation of monocot protoplasts. In: Galbraith, D.W., Bourque, D.P. and Bohnert, H.J. ed.. Methods in Cell biology: Methods in Plant Cell Biology. Part B, Academic Press, San Diego, 50: 383�399.

Maretzki, A. and Thom, M. (1987). UDP-glucose-dependent sucrose translocation in tonoplast vesicles from stalk tissue of sugarcane. Plant Physiol., 83 (2): 235�237.

Maretzki, A. and Thom, M. (1989). Vacuoles: isolation, purification, and uses. In: Bajaj, Y.P.S. ed. Biotechnology in Agriculture and Forestry: Plant protoplasts and genetic engineering II. Springer-Verlag, Berlin, 9: 459�479.


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326

Matsuoka, M., Terauchi, T., Kobayashi, M., Shoda, M. and Nakano, H. (1996). Isolation of protoplasts from suspension culture and subsequent shoot regeneration in sugarcane. J. Ag. Res., 30: 21�26.

Moore, P.H. (1987). Anatomy and morphology. In: Heinz, D.J ed. Sugarcane improvement through breeding. Elsevier, Amsterdam, 85�142.

Moore, P.H., Botha, F.C., Furbank, R.T. and Grof, C.P.L. (1997). Potential for overcoming physio-chemical limits to sucrose accumulation. In: Keating, B.A. and Wilson, J.R. ed. Intensive Sugarcane Production: Meeting the Challenges Beyond 2000. Proc. Sugar 2000 Symp., CAB Int., Wallingford, 141�156.

Potrykus, I., Harms, C. T., Lorz, H. and Thomas, E. (1977). Callus formation from stem protoplasts of corn (Zea mays L.). Mol. Gen. Genet., 156 (3): 347�350.

Srinivasan, C. and Vasil, I.K. (1986). Plant regeneration from protoplasts of sugarcane (Saccharum officinarum L.). J. Plant Physiol., 126: 41�48.

Taylor, P.W.J., Ko, H.-L., Adkins, S.W., Rathus, C. and Birch, R.G. (1992). Establishment of embryogenic callus and high protoplast yielding suspension cultures of sugarcane (Saccharum spp. hybrids). Plant Cell Tissue Organ Cult., 28: 69�78.

Thom, M. and Maretzki, A. (1985). Group translocation as a mechanism for sucrose transfer into vacuoles from sugarcane cells. Proc. Natl. Acad. Sci. USA, 82 (14): 4697�4701.

Thom, M., Maretzki, A. and Komor, E. (1982). Vacuoles from sugarcane suspension cultures�1. Isolation and partial characterisation. Plant Physiol., 69: 1315�1319.

Thom, M., Willenbrink, J. and Maretzki, A. (1983). Characteristics of ATPase from sugarcane protoplast and vacuole membranes [Saccharum spp. hybrid]. Plant Physiol., 58 (4): 497�504.

Vasil, I.K. and Vasil, V. (1980). Isolation and culture of protoplasts. Int. Rev. Cytol. Suppl., Academic Press, New York, 11B: 1�19.

Wagner, G.J. (1987). Isolation of mature vacuoles of higher plants: general principles, criteria for purity and integrity. Methods Enzymol., 148: 55�73.

Documents

ISOLATION OF PROTOPLASTS AND VACUOLES FROM … · Protoplast isolation from stem parenchyma cells The green peripheral part of the sugarcane stem (or culm), where vascular bundles