Plant Cell Reports

Plant Cell Reports (1987) 6:337-340

© Springer-Verlag 1987

Factors affecting protoplast electrofusion efficiency Lawrence J. Nea * and George W. Bates Department of Biological Science, Florida State University, Tallahassee, FL 32306-3015, USA Received November 12, 1986 Revised version received July 8, 1987 - Communicated by E. D. Earle

ABSTRACT The electrofuslon efficiency of protoplasts isolated from a carrot (Daucus carota) suspension culture was increased by treatment with 0.i mg/ml lysolecithin, 2.5% dimethylsulfoxide (DMSO), or 0.5 mM Ca 2+. The lysolecithin and DMSO treatments substantially increased protoplast lysis, whereas calcium treatment did not. The enzymes used for protoplast isolation were also found to have a dramatic effect on the efficiency of fusion. A mixture of Cellulysin and Driselase led to a two-fold enhancement of fusion as compared with Driselase alone. The stimulation by Cellulysin appears to be due to enzymatic modification of the cell surface. However, comparison of the time course for wall digestion with the development of susceptibility to electrofusion suggests that the effect of Cellulysin is not simply due to removal of the cell wall. Brief treatment of the cells with pronase or proteinase K also doubled the efficiency of fusion. Taken together, these results indicate that electrofusion efficiency can be enhanced by the method used for protoplast isolation; they also suggest that modification of membrane/cell-surface proteins during protoplast isolation may be particularly important in determining electrofusion efficiencies. Abbreviations: a.c., alternating current; d.c., direct current; DMSO, dimethylsulfoxide; NAA, naphthaleneacetic acid; PEG, polyethylene glycol INTRODUCTION Electrofusion is being applied increasingly to studies of somatic hybridization in plants (Bates et al., 1987). This technique, originally described by Senda et al. (1979) and later refined by Zimmermann and coworkers (Zimmermann, 1982), provides a rapid, simple procedure for fusing protoplasts.

Although electrofusion is usually described as reproducible and highly efficient, examination of the literature reveals that fusion efficiencies may range from a few percent to more than 60% (Bates et al., 1987; Tempelaar and Jones, 1985). As reported by Tempelaar and Jones (1985), electrofusion efficiency appears to differ for protoplasts isolated from different species and tissues. Similar observations have been made for the induction of protoplast fusion by PEG (Kao, 1981). Because of the desirability of obtaining efficient, reproducible fusions, we have studied the effects of various chemical treatments as agents for promoting electrofusion. Protoplasts from a carrot suspension culture were used as a model system. We found that the types of enzymes used for protoplast isolation can have a profound effect on the fusion efficiency. Our observations also suggest that the modification of membrane surface proteins plays an important role in protoplast electrofusion. MATERIALS AND METHODS A non-morphogenic (and non-fusogenic) suspension culture of Daucus carota (kindly provided by Dr. Wendy Boss, Dept. of Botany, North Carolina State University, Raleigh, N.C., U.S.A.) was maintained on ER medium (Eriksson, 1965) containing 1 mg/l NAA and 0.02 mg/l kinetin, on a rotary shaker at 125 r.p.m., under constant light (5 ~E/m2s). The culture was transferred weekly at a dilution of 1:10. Three to 5 days after subculturing, protoplasts were released by treating the cells with 2% Driselase ~n 0.4 M mannitol (adjusted to pH 5 . 1 with NaOH) for 5 hours. The digest was passed through a 62-~m mesh and centrifuged (130 x g) for 3 min. The protoplast pellet was washed three times with 10 ml of 0.4 M mannitol and resuspended in 2-3 ml of.0.4 M mannitol. The protoplasts were counted with a hemacytometer. For fusion, the protoplast density was adjusted to 5 x 104 cells/ml, and 10-15 ~i of protoplast suspension were introduced into a

* Present address: Petoseed Co., Research Center, Route 4, Box 1255, Woodland, CA 95695, USA Offprint requests to: G.W. Bates

338 D.E.P. Systems (Metamora, Mich., U.S.A.) Open Fusion Slide #2050S. This fusion chamber consists of a pair of parallel p l a t i n u m plates m o u n t e d (0.5 mm apart) on a glass microscope slide. So that fusion could be monitored, the chamber was m o u n t e d on the stage of an inverted microscope. The electric fields used for fusion were g e n e r a t e d by a Zimmermann Cell Fusion Power Supply (G.C.A. Precision Inc., Chicago, Ill., U.S.A.). Cell-to-cell contacts were induced by d i e l e c t r o p h o r e s i s with an a.c. field (160-200 V/cm, 600 kHz). After 2 min to allow for alignment of the protoplasts, the a.c. field strength was increased to 200-250 V/cm. Fusion was then induced by application of two d.c. pulses (600-1000 V/cm, 50 ~s in duration). Following fusion, the a.c. field was reduced to 30 V/cm. This weak field relieved the tension on the protoplasts, allowing them to coalesce, but p r e v e n t e d the protoplasts from d r i f t i n g out of the plane of focus. Fusion products were identified visually by m i c r o s c o p i c observation (400 X magnification) immediately after d e l i v e r y of the d.c. pulses. Fused protoplasts were recognized by the appearance of a distinct cytoplasmic connection between protoplasts in contact. The fusion e f f i c i e n c y was calculated from the fraction of protoplasts observed after fusion. Only those protoplasts that were aligned into chains by the a.c. field were counted; unaligned protoplasts were ignored. This p r o c e d u r e permits an accurate estimate of the response of the cells to the d.c. field alone. For the lysolecithin treatment, 0.5 ml of packed protoplasts were r e s u s p e n d e d in 1 ml of 0.i or 0.2 mg/ml lysolecithin in 0.4 M mannitol. After 5 min at room temperature, the protoplasts were washed three times with m a n n i t o l to remove the lysolecithin from the m e d i u m prior to fusion. In the case of the DMSO and CaCI 2 treatments, freshly isolated p r o t o p l a s t s were washed and then resuspended in 0.4 M mannitol containing DMSO (2.5-5%) or CaCI 2 (0.5 mM). DMSO and CaCI 2 were left in the m e d i u m during electrofusion. For C e l l u l y s i n treatments the protoplasts were isolated in a mixture of 0.5% Cellulysin and 1.5% Driselase. Protease treatments were applied to freshly isolated protoplasts as a 5-min treatment with 1 mg/ml pronase or a 3-min treatment with 0.25 mg/ml p r o t e i n a s e K in 0.4 M m a n n i t o l plus 0.5 mM MES, pH 5.5. The protoplasts were washed with mannitol prior to fusion. In order to v i s u a l i z e the cell wall, calcofluor (Phorwhite BBH, Mobay Chemical Co., Rock Hill, S.C.) was used at a concentration of 0.1 mg/ml in 0.4 M mannitol. After five minutes in calcofluor, the cells (or protoplasts) were washed twice with 0.4 M m a n n i t o l and o b s e r v e d by fluorescence m i c r o s c o p y with a Nikon V filter package. For culture, p r o t o p l a s t s were resus~ pended at a density of 5 x 104 cells/ml in ER m e d i u m containing 0.2 M mannitol, 1 mg/l NAA, and 0.02 mg/l kinetin and were grown in the dark at 27°C.

RESULTS AND D I S C U S S I O N E n h a n c e m e n t of E l e c t r o f u s i o n by Lysolecithin, DMSO, and Calcium. In initial experiments, we e x a m i n e d the effects of a number of compounds on the e l e c t r o f u s i o n of carrot protoplasts. DMSO, which has been used in combination with PEG to induce protoplast fusion (Menczel and Wolfe, 1984), significantly stimulated e l e c t r o f u s i o n (Fig. I). The stimulation, although small, is statistically significant. However, DMSO increased cell lysis substantially. This effect increased with increasing DMSO concentrations. For example, treatment with 5% DMSO resulted in the lysis of more than half the protoplasts. Lysis of DMSO treated protoplasts was o b s e r v e d during both protoplast washing and electrofusion. Although omitting the final wash improved the situation, the remaining degree of lysis probably offsets any advantages of this compound. The greater p r o t o p l a s t fragility observed here than when DMSO is used in combination with PEG (Menczel and Wolfe, 1984) may reflect the greater physical forces exerted on the m e m b r a n e during electrofusion.

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~5%

Figure i. Effects of small m o l e c u l e s and Cellulys[n on e l e c t r o f u s i o n of carrot protoplasts. Protoplasts were isolated in 0.4 M m a n n i t o l with 2% Driselase, washed, and then treated for 5 min with 0.i mg/ml lysolecithin, 2.5% DMSO, or 0.5 mM CaCI 2. Lysolec i t h i n - t r e a t e d protoplasts were washed prior to electrofusion, but DMSO and CaCI 2 were left in the m e d i u m (0.4 M mannitol) during fusion. The Cellulysin treatment (+) consisted of protoplasts isolated with 1.5% D r i s e l a s e plus 0.5% Cellulysin. The control (-) c o n s i s t e d of protoplasts isolated with 2% D r i s e l a s e alone. The data p r e s e n t e d are average values of 7-9 d e t e r m i n a t i o n s ! the S.E. Differences in fusion b e t w e e n treated protoplasts and their respective controls are s t a t i s t i c a l l y significant (p < 0.05).

Lysolecithin was also found to stimul a t e the e l e c t r o f u s i o n of carrot p r o t o p l a s t s (Fig. i). This compound can by itself induce fusion in some animal cell systems. However, lysolecithin has not been found to induce the fusion of plant protoplasts either when used alone or in c o m b i n a t i o n with PEG (Constabel and Kao, 1974; Gleba and

339 Sytnik, 1984). In fact Constabel and Kao (1974) found that lysolecithin treatment reduced the efficiency of PEG-induced fusion. As with DMSO, lysolecithin treatment led to protoplast lysis. However, the degree of lysis was less and could be controlled by washing the cells before electrofusion. Lysis in response to lysolecithin has been observed before (Constabel and Kao, 1974; Gleba and Sytnik, 1984). Although we did not determine plating efficiencies, the lysolecithin-treated carrot protoplasts readily formed callus when cultured. Of all the small molecules tested, Ca 2+ shows the greatest promise as a useful agent for enhancing electrofusion. Treatment of the protoplasts with 0.5 mM CaCl 2 significantly enhanced fusion (Fig. i) without inducing lysis. Concentrations above 0.5 mM could not be tested because they interfered with a.c.-field-induced cell alignment. A beneficial role of Ca 2+ in protoplast fusion has often been observed (Gleba and Sytnik, 1984) and can probably be attributed to calcium's ability to neutralize surface charges and reduce membrane fluidity. Spangenberg and Schweiger (1986) and Tempelaar et al. (1987) also report that protoplast electrofusion is enhanced by CaCI 2 and Watts and King (1984) found that Ca 2+ reduced the sensitivity of mesophyll protoplasts to electric-field-induced lysis. Although field-induced lysis of carrot suspensioncell protoplasts was not affected by Ca 2+, this difference may simply reflect the greater stability of suspension-cell protoplasts than mesophyll protoplasts. Enhancement of Fusion by Cellulysin and Proteases. Comparison of the controls for the different experiments shown in Fig. 1 indicates a considerable amount of day-to-day variation in the extent of fusion. Fusion efficiency was essentially constant from day 2 to day 6 (data not shown). Thus, variation did not appear to reflect the use of protoplasts isolated on different days of the suspension culture's subculture interval. While trying to determine the source of the day-to-day variation in fusion efficiency, we observed that the inclusion of Cellulysin with Driselase during protoplast isolation dramatically increased s u b s e q u e n t electrofusion (Fig. i). This enhancement of fusion by Cellulysin was unexpected because Driselase alone caused almost complete conversion of the carrot suspension culture cells to a clean and uniform protoplast preparation. Use of 2% Cellulysin (without Driselase) did not speed protoplast release, but it did cause extensive clumping of the protoplasts and possibly spontaneous fusion as well. The use of boiled Cellulysin, in combination with Driselase (unboiled), for protoplast isolation did not enhance electrofusion (Table i). However, dialyzed Cellulysin was fully active. Thus, the enhancement of fusion by Cellulysin seems to be due to an enzyme present in Cellulysin rather than to a fusogenic, low-molecularwelght contaminant, such as has been implicated in the induction of fusion by PEG (Honda et al., 1981).

Table i. Effects of boiled or dialyzed Cellulysin on electrofusion. Carrot protoplasts were isolated by a 5-h digestion in 2% Driselase (control) or by a 5-h digestion in 1.5% Driselase plus 0.5% Cellulysin that had been either boiled (5 min), dialyzed, or left untreated. The protoplasts were purified and subjected to electrofusion. The data are presented as the average percentage of fusion ~ S.E. Each treatment was repeated four times. Treatment

% Fusion

Control Cellulysin Boiled Dialyzed

16.1 37.3 15.3 37.2

~ ~ ~ ~

0.7 0.7 0.9 3.1

The enhancement of fusion by Cellulysin could be the result of incomplete wall removal by Driselase. To test this hypothesis, we used calcofluor staining to monitor wall digestion. After 2.5 hours of digestion with Driselase, most of the cells were spherical and would not stain with calcofluor. Despite the apparently complete removal of their cell walls, these spherical, nonstaining cells were resistant to fusion. Time courses for wall digestion and the development of susceptibility to electrofusion are shown in Table 2. Electrofusion efficiencies remained low after 2.5 hours of digestion even though almost all the cells had been converted to protoplasts. Extending the digestion to 4 hours resulted in improved fusion. The lack of correlation between .these time courses suggests that residual cell-wall material is not limiting the fusion of protoplasts isolated with Driselase. This conclusion rests on the sensitivity of calcofluor for assessing wall digestion. Admittedly, calcofluor only binds to one fraction of the cell wall--cellulose; thus the possibility that Cellulysin removes other wall polysaccharides more effectively than does Table 2. Relationship between the extent of wall digestion and the efficiency of electrofusion. Carrot suspension cells were digested with 2% Driselase for the periods indicated; protoplasts were then isolated and subjected to electrofusion. Data are the averages of 4 trials ~ S.E. The percentage of digestion was determined from the fraction of cells retaining calcofluor fluorescence after the period indicated. Only nonfluorescent cells (protoplasts) were scored for fusion. Length of Digestion 1.0 h 2.5 h 4.0 h

% Fusion

% Digestion

2.4 ~ 0.7 4.7 ~ 0.8 ii.3 ~ 1.6

20-25 > 95 > 99

340 Driselase cannot be eliminated. Nonetheless, these observations suggest that the enhancement of fusion by Cellulysin results from some enzyme activity other than a polysaccharidase in this enzyme mixture. Driselase and Cellulysin are known to contain proteases and lipases (Vasil and Vasil, 1980). We considered it unlikely that lipase activity in Cellulysin was responsible for the enhancement of electrofusion because even prolonged digestions in Cellulysin (2% for 5 h) did not lead to cell lysis. However, treating protoplasts isolated with Driselase briefly with proteases mimicked the effect of Cellulysin. Thus, a 5-min treatment with 1 mg/ml pronase or 3 min with 0.25 mg/ml proteinase K doubled the efficiency of electrofusion (from 6.9% to 13.8% for pronase and from 7.4% to 16.6% for proteinase K). These observations suggest that the enhancement of the electrofusion of carrot protoplasts by Cellulysin (as compared w i t h p r o t o p l a s t s isolated with Driselase alone) could be due to proteases in this enzyme mixture. Proteases have previously been observed to enhance fusion. Constabel and Kao (1974) found that the PEG-induced fusion of pea and Vicia protoplasts by PEG was reduced by trypsin and papain but it was stimulated by lysozyme. Kameya (1979) has reported that pronase enhances dextran-induced protoplast fusion. Pronase is also widely reported to stimulate and even to be required for the electrofusion of animal cells (Zimmermann, 1982). On the other hand, Spangenberg and Schweiger (1986) found that the electrofusion of Brassica napus hypocotyl protoplasts was not affected by treatment with proteinase K, although fusion was stimulated by dispase. Ruzin and McCarthy (1986) report that trypsin, pronase and dispase all facilitate the electrofusion of tobacco mesophyll protoplasts; however, dispase was the least effective. Our experiments show that pronase and proteinase K can be used to enhance the electrofusion of carrot suspension-cell protoplasts. However, the practical value of such a treatment is unclear. Based on their ability to exclude 0.1% Evans blue, we found that carrot protoplasts remain viable after a 5-min treatment with 1 mg/ml pronase (24 h after isolation and treatment, viability ranged from 79% to 86% for both pronase-treated and untreated protoplasts). However, a 10-min treatment with pronase proved to be toxic. The day-to-day variability of electrofusion efficiency that we ~ b s e r v e d remains unexplained. It must be due to some change in the properties of the cell membrane or cell wall that we have been unable to control. The enhancement of protoplast elec-

trofusion by proteases points to membrane proteins as one cellular component that may be important in determining the efficiency of fusion. It is also conceivable that changes in membrane-surface proteins is a factor leading to day-to-day variations in fusion efficiency. Certainly membrane proteins would differ between species and cell types. Such differences might explain why other laboratories have found that certain proteases are effective for enchancing the fusion of some protoplast preparations but not others. Whether or not the stripping away of membrane proteins proves to be important in prot0plast fusion, our work indicates that the enzymes chosen for protoplast isolation must be optimized for the efficiency of fusion as well as for protoplast release for each new cell system.

Acknowledgment. This work was supported by U.S. Department of Agriculture competitive research grant 83-CRCR-I-1257 to G.W.B.

REFERENCES Bates GW, Nea LJ, Hasenkampf, CA (1987) In: Sowers AE (ed) Cell Fusion, Plenum Press, New York, pp. 479-496 Constabel F, Kao KN (1974) Can J Bot 52:1603- 1606 Eriksson, T (1965) Physiol Plant 18:976-993 Gleba YY, Sytnik KM (1984) Protoplast Fusion, Genetic Engineering in Higher Plants. Springer-Verlag, Berlin Honda K, Maeda Y, Sasakawa S, Ohno H, Tsuchida E (1981) Biochem Biophys Res Comm 101:165-171 Kameya T (1979) Cytologia 44:449-456 Kao KN (1981) In: Han H (ed) Plant Tissue Culture, Proceedings of the Beijing Symp, Pitman International Series in Applied Biology, Pitman, London, pp 331-339 Menczel L, Wolfe K (1984) P1 Cell Reports 3:196-198 Ruzin SE, McCarthy SC (1986) P1 Cell Reports 5:342-345 Senda M, Takeda J, Abe S, Nakamura T (1979) Plant Cell Physiol 20:1441-1443 Spangenberg G, Schweiger H-G (1986) Eur J Cell Biol 41:51-56 Tempelaar MJ, Jones MGK (1985) Planta 165:205-216 Tempelaar MJ, Duyst A, DeVlas Sy, Krol G, Symmonds C, Jones MGK (1987) Plant Sci 48:99-105 Vasil IK, Vasil V (1980) Int Rev Cytol, Suppl lIB:I-17 Watts JW, King JM (1984) Biosc Rep 4:335-342 Zimmermann U (1982) Biochim Biophys Acta 694:227-277