Generation and isolation of cyanobacterial inside-out thylakoid vesicles

ARCHIVES

Vol.

OF BIOCHEMISTRY

AND

265, No. 2, September,

Generation

BIOPHYSICS

pp. 321-328,1988

and Isolation of Cyanobacterial

FREDRIK

NILSSON,* AND

*Department tDepa,rtment

of Biochemistry, of Physiology, and SDepartment

Received

Inside-out

Thylakoid

DAVID J. SIMPSON,t ALISON BERTIL ANDERSSON*s’

Vesicles

C. STEWART,+

Arrhenius Laboratories, University of Stockholm, S-106 91 Stockholm, Sweden; Carlsberg Laboratory, Gl, Carl&erg Vej 10, DK-2500 Copenhagen, Denmark; of Biochemistry, University of Cambridge, Tennis Court Road, Cambridge CB2 l&W, United Kingdom September

23,198’7,

and in revised

form

May

2, 1988

A method has been designed to prepare inside-out thylakoid vesicles from a cyanobacterial splecies (Phormidium laminosum). Everted thylakoid vesicles could be generated by Yeda press treatment after induced membrane pairing. Membrane pairing was induced either by addition of high concentrations of M$+ ions or by lowering the pH of the fragmentation media. The inside-out vesicles were isolated by aqueous polymer two-phase partition. The membrane orientation was determined by proton translocation studies and freeze-fracture electron microscopy. o 1988 Academic press, I,,~.

Cyanobacteria are free-living procaryotes capable of oxygenic photosynthesis, requiring both photosystem I and photosystem II (1). Although their photosynthetic apparatus shows many similarities with that of eucaryotic cells there are also some important differences. These include an extrinsic light harvesting system (the phycobilisomes), lack of chlorophyll b (2), simpler subunit structure of photosystem II (3), alternative electron carriers (4), unstacked thylakoids (5), and a low photosystem II/I ratio (6). Moreover, the cyanobacterial thylakoid membrane is thought to carry thle components necessary for respiration (‘7). Cyanobacteria as a model system for studying the oxygenic photosynthesis has gained increasing attention since they,, as procaryotic organisms, can be genetically transformed (8). Our understanding of the molecular organization of the cyanobacterial energy transducting system, apart from the phycobilisomes, is mainly based upon assumed analogies with higher plant thylakoids. The organization of the cyanobac1 To whom

correspondence

should

terial thylakoid membrane is poorly characterized, particulary in respect to its transverse asymmetry. This stands in contrast to the situation with the plant thylakoid membrane, where studies on the transverse asymmetry have been made possible by the development of a method to obtain inside-out thylakoid vesicles (9-11). No such preparation method has so far been available for cyanobacterial thylakoids. Inside-out thylakoid vesicles from higher plant chloroplasts are generated by mechanical fragmentation and isolated by aqueous polymer two-phase partition (9-11). This separation technique, which has been applied to a number of biological compounds, separates membranes according to differences in surface properties (12). In this communication we present a method to generate and isolate inside-out vesicles from the photosynthetic membrane of the cyanobacteria Phormidium laminosum. MATERIAL

AND

METHODS

Polyethylene glycol was obtained 3350 from Union Carbide (New York,

be addressed. 321

0003-9861/88 Copyright All rights

as Carbowax NY). Dextran

$3.00

0 1988 by Academic Press, Inc. of reproduction in any form reserved.

322

NILSSON

T-500 was obtained from Pharmacia Fine Chemicals (Uppsala, Sweden). Thylakoid preparation P. laminosum was grown on a low salt medium (13) in white light at 45’C according to (14). Thylakoids were prepared by lysozyme treatment followed by centrifugation as described in (14). The thylakoid preparation was stored in liquid nitrogen. The bulk of the phycobilisomes was removed by repeated washing in a high salt buffer containing 10 mM sodium phosphate, pH 7.4, 250 mM NaCl, and 10% glycerol by centrifugation at 10,OOOg for 10 min. Fragmentation procedure. Three different media were used for fragmentation: (a) 10 mM Tricine,’ pH 7.4,lOO mM sucrose, 10% glycerol; (b) 10 mM Tricine, pH 7.4, 100 mM sucrose, 75 pM LaCI,, 10% glycerol; and (c) 10 mM Tricine, pH 7.4,lOO mM sucrose, 50 mM MgClz, 10% glycerol. Cyanobacterial thylakoids (approx 0.6 mg chlorophyll/ml) were suspended in one of the three media and passed twice through a Yeda press operated at a nitrogen pressure of 15 MPa. When medium (a) was used the thylakoids were first kept at pH 7.4 and immediately prior to fragmentation the pH was lowered to pH 4.7 by addition of dilute HCl. Immediately after the Yeda press treatment the pH was adjusted back to 7.4 by addition of NaOH. The three Yeda press homogenates were centrifuged at 40,OOOg for 30 min and washed twice in 10 mM sodium phosphate, pH 7.4, 5 mM NaCl, 100 mM sucrose, 10% glycerol. The final pellets were suspended in this medium and passed twice more through the Yeda press at a pressure of 10 MPa. All preparation and fragmentation steps were performed at 4°C. Phase partition of thylakoid fragments, Phase partition was performed at 4°C in a 25 g two-phase system containing 5.9% (w/w) dextran T-500, 5.9% (w/w) polyethylene glycol 3350, 10 mM sodium phosphate buffer, pH 7.4, 5 mM NaCI, 40 mM sucrose, and 10% (w/w) glycerol. Material containing 10 mg chlorophyll was added to the system which was then mixed. The system was settled by low-speed centrifugation. The upper phase was collected and repartitioned with a pure lower phase to yield the T2 fraction. The original lower phase (Bl) was repartitioned twice with pure upper phase to yield the B3 fraction. The B3 and T2 fractions were collected by centrifugation at 100,OOOg for 45 min after twofold dilution in 10 mM sodium phosphate, pH 7.4, 5 mM NaCI, 100 mM ’ Abbreviations used: Tricine, N-[2-hydroxy-l,lbis(hydroxymethyl)ethyl]glycine; Hepes, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid; Mes, 4morpholineethanesulfonic acid; EF, endoplasmic face; PF. protoplasmic face; SDS, sodium dodecyl sulfate; PS, photosystem; DCIP, 2,6-Dichlorophenol indophenol; DCMU, 3(3,4-Dichlorophenyl)-l,l-dimethylurea.

ET

AL.

sucrose, 10% glycerol. For proton translocation measurements the pellets were suspended in 40 mM KCl, 10% glycerol. Viscosity problems restricted the use of 20-30% glycerol which is optimal for preserving the photosystem II activity of the cyanobacterial thylakoids (15). Analyses. Chlorophyll a was estimated by absorbance measurements at 680 nm or according to Arnon et al. (16). Light-induced proton translocation studies were performed using a glass electrode. Vesicles containing 20 fig chlorophyll were suspended in 1 ml of a medium containing 40 mM KCl, 10% glycerol, 1 mM phenyl-p-benzoquinone at 25°C. Photosystem II activity was assayed in a Clarktype oxygen electrode. Thylakoid vesicles (20 pg chlorophyll) were suspended in 1 ml medium containing 10 mM Hepes-NaOH, pH 7.0, 10 mM MgCIz, 25% (w/w) glycerol, 1 mM phenyl-p-benzoquinone. Photosystem I activity was assayed by measurements of oxygen uptake. Thylakoid vesicles (20 rg chlorophyll) were suspended in a medium containing 120 FM methylviologen, 0.6 mM NaN3, 80 pM DCIP, 32 mM Na-ascorbate, 10 pM DCMU, 40 mM sodium phosphate, pH 7.0, 1 mM NaCl, and 5 mM CHaNHZ. Light from two opposite placed 375 W tungsten lamps supplied with red filters was used for excitation. Freeze-fracture. Vesicle preparations suspended in 10 mM Mes, pH 6.5, and 5 mM MgClz (at about 1 mg chlorophyll a/ml) were made 30% with respect to glycerol and handled at 4°C. They were frozen in small aliquots on gold specimen disks by immersing into Freon 22 cooled by liquid nitrogen, and stored under liquid nitrogen until used for freeze-fracturing. Specimens were fractured at -110°C and rotary shadowed at an angle of 22.5” with Pt/C from an electron beam evaporation gun to a depth of 20 A. The replicated surface was coated with about 300 A of carbon and cleaned for 2 h in concentrated sodium hypochlorite solution, followed by dilute chromsulfuric acid overnight. Replicas were washed four times in distilled water and transferred to flamed 200 mesh copper grids. They were examined in a Siemens Elmiskop 102 operated at 80 kV and calibrated at 50,000X and 100,000X magnification against catalase crystals (17). RESULTS

AND

DISCUSSION

According to the mechanism of formation for inside-out vesicles from higher plant thylakoids, the membranes are required to be in a tightly appressed state during the mechanical disintegration (10, 18). Cyanobacterial thylakoids do not have stacks of appressed membranes characteristic of plant thylakoids. Therefore, in

CYANOBACTERIAL

I

THYLAKOID

r

PHYCOBILISOME

I

INSIDE-OUT

PREPARATION

I

1 REMOVAL

YEDA-PRESS

x 2

(15

HPa)

I

-I-

I1

TRANSFER CENTRIFLJGATION

r1YEDA-PRESS

TO NEW BUFFER x 2 (40 000

1

I I

PHASE

FIG. 1. Scheme for thylakoid fragmentation.

x q)

1

x 2 (10

MPa)

PARTITION

J

I

I

phycobilisome

removal

and

order to generate inside-out vesicles from cyanobacteria, membrane appression has to be induced by manipulations of the membrane and its surrounding milieu. It was reasoned that two major obstacles might prevent tight membrane pairing of the cyanobacterial thylakoids: the presence of bulky protruding phycobilisomes on the outer thylakoid surface and

t

THYLAKOID

83

T2

rise

DARK

I fpLIGHT

323

repulsive intermembrane electrostatic forces. Therefore the phycobilisomes were removed by repeated washing in a high salt buffer. The electrostatic repulsion between the thylakoid membranes was reduced by neutralization or screening of surface negative charges. Neutralization was achieved by adjusting the pH to 4.7, which is the isoelectric point of plant thylakoid membranes (19) and was assumed to be applicable also in the case of the stripped cyanobacterial thylakoids. This method of low pH treatment in combination with Yeda press disruption has been successfully applied to obtain functional inside-out vesicles from destacked and randomized higher plant thylakoid membranes (10). Phase partition of the low pH-treated and fragmented cyanobacterial thylakoids (Fig. 1) resolved two membrane populations, one major partitioning to the upper phase and one minor partitioning to the lower phase. After the subsequent partition steps the T2 fraction contained 80% of the material on a chlorophyll basis whereas the B3 fraction contained only 2% of the material. As an analysis of the membrane orientation of the thylakoid vesicles, proton translocation was measured. Phenyl-pbenzoquinone was used as electron acceptor. Previous studies on higher plant thylakoid vesicles have shown that phenyl-pbenzoquinone mediates electron transport in a photosystem II reaction which leads to an uncoupler-sensitive proton translocation across the membrane (20, 21). The

LIGHT

pH

VESICLES

DARK

FIG. 2. Light-induced photosystem II-mediated proton translocation of cyanobacterial thylakoid vesicles ‘obtained by phase partition. The height of the arrow represents 10 nmol H+. The extent of proton uptake or extrusion was determined by addition of 10 nmol NaOH. White arrows show when the light is switched on, dark arrows show when the light is turned off.

324

NILSSON

proton translocation studies on the T2 and B3 fractions revealed that the vesicles of the T2 fraction had a normal light-induced reversible proton uptake (Fig. 2). In contrast, under the same conditions the B3 fraction showed a light-induced reversible proton extrusion. These results show that the B3 fraction is dominated by vesicles with a reversed proton translocation, suggesting that these vesicles are turned inside-out with respect to the normal orientation of the thylakoid membrane. These results are analogous to results obtained from similar subfractionation studies on higher plant thylakoids (10). An independent way to determine the orientation of membrane vesicles is freeze-fracture electron microscopy (18). By analyzing the curvature of the endoplasmic face (EF) and protoplasmic face (PF) the sidedness of single vesicles in a membrane population can be determined accurately. Thylakoid vesicles of normal sidedness show convex EF and concave PF whereas everted vesicles show concave EF and convex PF. The two fracture faces of unfragmented Phormidium thylakoids could be distinguished and identified from their morphology. The EF face contained fewer freeze-fracture particles per unit area than the PF face, and the particles were slightly larger (9.5 vs 7.0 nm) in size (Fig. 3). There was no indication of particle alignment on either face, and the main difference between E and P faces was thus particle number per unit area. The freezefracture appearance of P. laminosum thylakoids resembled that of the blue green algae Anacystis nidulans (22) and Anabaena cylindrica (23), in which EF particles are randomly distributed. Both the B3 and T2 fractions contained spherical and unilamellar vesicles of uniform size (80 and 120 nm in diameter) (Fig. 4). The vesicle size was estimated at magnifications which had been calibrated with diffraction grating replicas, and by measuring only those vesicles which had been cleaved close to their center (as judged from the height and depth of the vesicle). This will lead to a slight underestimation of both minimum and maximum diameters.

ET

AL.

Because of their similar appearances, it was not always possible to discriminate between EF and PF faces in concave vesicles, unless they were very shallow. A number of micrographs were analyzed for types of vesicles present and the results are given in Table I. In most cases, only convex vesicles were counted. The same analysis was applied to micrographs from the T2 fraction (Fig. 5). While the latter contained equal proportions of normal and everted vesicles, the B3 fraction was enriched (77%) in vesicles of everted sidedness (Table I). This 3:l enrichment of inside-out vesicles of the B3 fraction is similar to that previously shown for spinach thylakoid vesicles (18). In contrast to the spinach T2 fraction, which was greatly enriched in right-side-out vesicles (8:1), that from Phormidium contained equal proportions of right-side-out vesicles and inside-out vesicles (Table I). Still the T2 fractions showed a net proton uptake, which can be explained by the fact that the

FIG. Phormidium

fracture number X108.800.

3. Freeze-fracture electron micrograph of thylakoids showing the two different faces (EF and PF). These differ mainly in of particles per unit area. Magnification

CYANOBACTERIAL

INSIDE-OUT

THYLAKOID

VESICLES

325

FIG. 4. Freeze-fracture appearance of vesicles in the B3 fraction after aqueous phase partition. Inside-out vesicles are convex PF (asterisks) and concave EF (small arrowheads), whereas those of normal sidedness are concave PF (larger arrowheads) and convex EF (arrow). Magnification X57,800. FIG. 5,. Freeze-fracture vesicles from the T2 fraction, labeled as in Fig. 4. Magnification X57,800.

right-side-out vesicles retain more of their photochemical activities compared to everted vesicles (Table II).

We also tried to obtain inside-out vesicles by Yeda press treatment of thylakoids after addition of 75 FM La3+ ions. Such tri-

326

NILSSON TABLE

I

DETERMINATION OF SIDEDNESS OF VESICLES FROM PARTITIONED CYANOBACTERIAL THYLAKOIDS BY FREEZE-FRACTURE ELECTRON MICROSCOPY

Fraction T2 B3

Convex EF + concave PF (%I

Concave EF + convex PF (%o)

50 23

50 77

Total counted 101 52

valent ions are known to bind to and neutralize negative surface charges (24). However, after phase partition of such membrane fragments virtually no material was recovered in the lower phase, indicating that no inside-out vesicles could be isolated. Moreover, the membrane vesicles were inactive in both oxygen evolution and proton translocation. La3+ ions obviously cause irreversible damage to the membrane structure and function and could not be used to obtain active everted cyanobacterial thylakoids. Recently it has been shown that La3+ ions cause damage to the inner thylakoid surface of plant thylakoids (25). As an alternative to charge neutralization, shielding of surface negative charges by addition of divalent ions such as Mga can induce membrane pairing at neutral pH (24). For higher plant thylakoids only 2-3 mM Mga+ is required to form tightly appressed grana stacks required for the formation of inside-out vesicles. When such a low Me concentration was used in the press medium for cyanobacterial thylakoids no B3 material could be obtained (not shown). A Mgz+ concentration of 50 mM prior to fragmentation was required to give vesicles with a partition behavior resembling that obtained after Yeda pressing of the thylakoids at low pH. Thus around 80% of the material was recovered in the T2 fractions, whereas some 2% was recovered in the B3 fraction. The proton translocation studies (Fig. 2) revealed a light-induced proton uptake and extrusion for the T2 and B3 vesicles, respectively. Thus 50 mM Me is also able to induce membrane pairing of the cyanobacterial thylakoid membrane and allow inside-out

ET AL.

vesicles to be formed. It should be stressed that the moderately high M$+ concentration did not cause much damage to the photochemical activities. No freeze-fraeture studies were performed on these vesicles but their identical partition and proton translocation behavior as compared to those vesicles obtained from the low pH treatment suggest a similar sidedness distribution. High concentrations of Mg+ ions have previously been used to create membrane TABLE

II

PHOTOSYSTEM I AND PHOTOSYSTEM II ACTIVITY OF FRACTIONS OBTAINED BY YEDA PRESS TREATMENT AND PHASE PARTITION OF CYANOBACTERIAL THYLAKOIDS Electron transport rates

Fraction Untreated thylakoids Phycobilisome-free thylakoids H+-treated vesicles After first fragmentation After second fragmentation T2 fraction B3 fraction B3 fraction + spinach plastocyanin Me-treated vesicles After first fragmentation After second fragmentation T2 fraction B3 fraction

PSI (%I

PS II (o/o)

100 89

100 84

81 60 57 34

71 61 49 23

46 75 60 65 35

77 71 50 28

Note. Fractions were obtained as shown in Fig. 1. Photosystem II activity was assayed by measurement of oxygen evolution with a Clark oxygen electrode in an assay medium containing 1 mM phenyl-pbenzoquinone, 25% glycerol, 10 mM MgCla, 10 mM Hepes-NaOH, pH 7.0. Photosystem I activity was assayed by measurements of oxygen uptake in a medium containing 0.6 mM NaNa, 120 pM methylviologen, 80 pM DCIP, 32 mM Na-ascorbate, 10 PM DCMU, 40 mM sodium phosphate, pH 7.0, 1 mM NaCl, and 5 mM CH&IHa. Total volume, 1 ml. The specific activity in the untreated thylakoid preparation was 490 for the PS II measurements and 520 for the PS I measurements, expressed as micromoles Oa per milligram chlorophyll per hour. Electron transport rates are expressed relative (%) to that of the untreated thylakoids.

CYANOBACTERIAL

INSIDE-OUT

pairing in a chlorophyll b-less barley mutant in order to generate inside-out vesicles (26). Such a relatively high Me concentration may be a general requirement to induce tight membrane pairing at neutral pH in thylakoids devoid of the lightharvesting chlorophyll a/b complex of photosystern II. The photochemical activity of the Phormidium thylakoid membrane was followed throughout the various preparation steps (Table II). Both methods of preparation (Me and H+ incubation) resulted in some loss of photosystem I and photosystern II activity. Some 60-75% of the control activity (photosystem I or photosystern II) remained after the fragmentation. During phase partition there was further loss of photochemical activity, particularly for th[e B3 fractions (Table II). It is not likely that this is due to any deleterious action by the polymers since their presence has been shown to be protective rather than destructive for a number of biological materials (27). Rather, the decline of photochemical activity in the B3 fractions should be due mainly to the release of extrinsic proteins from the exposed inner thylakoid surface of the everted vesicles. In the case of photosystern I a release of plastocyanin is a likely reason for loss of activity (28). This assumption is supported by the fact that addition of spinach plastocyanin partly restores the photosystem I activity. We have previously shown that cyanobacteria contain a 9-kDa photosystem II protein which is easily lost from cyanobacterial thylakoid particles under conditions of low glycerol concentration (3, 15). This probably accounts for the relatively low photosystem II activity of the B3 fraction. Preliminary studies on the T2 and B3 material by SDS-polyacrylamide gel electrophoresis revealed no apparent differences in polypeptide composition between the two vesicle populations, indicating that theiir total protein composition should be similar. This indicates that the everted vesicles, despite a low yield, appear to be representative of the overall cyanobacterial membrane. Attempts were made to increase the

THYLAKOID

327

VESICLES

yield of inside-out vesicles by lowering the temperature at which the phase partition was performed. However, when the B3 fractition contained more than 2% of the total chlorophyll a content no reverse proton translocation could be detected, indicating an increased proportion of contaminating right-side-out vesicles. The freezefracture studies show that the total proportion of inside-out vesicles after fragmentation is about 50%. The main problem of obtaining higher yield of purified everted vesicles therefore seems to be the phase system step, which is not as selective in discriminating the sidedness of cyanobacterial thylakoid vesicles compared to higher plant thylakoid vesicles. At present we have no explanation for this discrepancy or what may be the difference between inside-out vesicles partitioning with the right-side-out vesicles to the upper phase and those having high affinity for the lower phase. Despite the rather low yield of inside-out vesicles and the reduced photochemical activity, these everted vesicles open up interesting experimental possibilities. The low yield can be compensated for by using large phase volumes, which can be done easily since polymer two-phase systems can be scaled up a hundredfold (12,29). ACKNOWLEDGMENT This work was supported Science Research Council.

by the Swedish

Natural

REFERENCES 1. PADAN, E. (1979) Annu. Rev. Plant. Physiol. 30, 27-40. 2. GANTT, E. (1981) Annu. Rev. Plant. Physiol 32, 327-342. 3. STEWART, A. C., LJUNGBERG, U., AKERLUND, H.-E., AND ANDERSSON, B. (1985) Biochim. Biophys. Acta 808, 353-362. 4. CROFTS, A. R., AND WOOD, P. M. (1978) in Current Topics in Bioenergetics (Sanadi, D. R., and Vernon, L. P., Eds.), Vol. 7, pp. 135-244, Academic Press, New York. 5. STAINER, R. Y., AND COHEN-BAZIRE, G. (1977) Anna Rev. Microbial. 31,225-274. 6. MIMURO, M., AND FUJITA, Y. (1977) Biochim. Biophys. Acta 459, 376-389. 7. BINDER, A. (1982) J. Bioenerg. Biomembr. 14, 271-286.

NILSSON

328

8. GRIGORIEVA, G., AND SHESTAKOV, S. (1982) FEMS MicrobioL Z&t. 13,367-370. 9. ANDERSSON, B., AND AKERLUND, H.-E. (1978) Biochim.

Biophys.

Acta 503,462-474.

10. ANDERSSON, B., SUNDBY, C., AND ALBERTSSON, P.-A. (1980) Biochim. Biophys. Acta 599, 391-402.

11. ANDERSSON, B., SUNDBY, C., AKERLUND, H.-E., AND ALBERTSSON, P.-A. (1985) Physiol. Plant. 65,322-330.

12. ALBERTSSON, P.-A. (1986) Partition of Cell Particles and Macromolecules, 3rd ed., Wiley, New York. 13. CASTENHOLZ, R. W. (1970) Schweiz 2. HydroL 32,

ET AL. 20. GOULD, J. M., AND IZAWA, S. (1974) Biochim. Biophys. Acta 333,509-524. 21. AKERLUND, H.-E., ANDERSSON, B., AND ALBERTSSON, P.-A. (1976) Biochim. Biophys. Acta 449, 525-535. 22. ARMOND, P. A., AND STAEHLIN, L. A. (1979) NutL Acad. Sci. USA 76, 1901-1905. 23.

chim.

AND

K. (1974)

WADA,

B&him.

Biophys.

18.

ANDERSSON,

B.,

HANSEN,

Cadsberg

G. (1978)

D.

Cadsberg

J.,

AND

26. 27.

BROOKS,

Cadsberg

HOYER-

Res. Commun.

43,

19.

AKERLUND, AND

H.-E.,

ANDERSSON,

ALBERTSSON,

Biophys.

Acta

P.-A.

552,238-246.

B., PERSSON,

(1979)

Biochim.

A.,

HIERHOLZER,

P. D.,

Res. Commun. D. in

E.,

SHARP,

IVEY,

S., AND

50,347-367. K.

A.,

AND

FISHER,

D.

(1985) Partitioning in Aqueous Two-Phase Systems (Walter, E., Brooks, D. E., and Fisher, D., Eds.), pp. 11-84, Academic Press, New York.

44,

77-89.

M.,

Physiol 33,

BERG, S. D. (1987) in Progress in Photosynthesis Research (Biggins, J., Ed.), Vol. 11, PP. 321-324, Martinus Nijhoff, Dordrecht. BASSI, R., HINZ, U., AND BARBATO, R. (1985)

28.

SIMPSON,

Bic-

L. A. (1979)

ZITKUS,

Acta

Res. Commun.

STAEHLIN,

Acta 546,373-382. (1982) Annu. Rev. Plant.

25.

357,231-245. 17. SIMPSON, D. J. (1979) 305-336.

AND

Biophys.

24. BARBER, J. 261-295.

538-551.

14. STEWART, A. C. AND BENDALL, D. (1980) Biochem. J. 188,351-361. 15. STEWART, A. C., SICZKOWSKI, M., AND LjaXcBERG, U. (1985) FEBS Lett. 193, 175-179. 16. ARNON, D. I., MCSWAIN, B. D., TSUJIMOTO, H. Y.,

T. H.,

GIDDINGS,

PTOC.

29.

HAEHNEL, B. (1981) HUDSTEDT, (1985)

W.,

BERZBORN,

Biochim. H.,

KRONER,

R. J., AND

Biophys. K. H.,

Ada AND

ANDERSSON,

637,389-399. KULA,

M.-R.

Partitioning in Aqueous Two-Phase Systems (Walter, E., Brooks, D. E., and Fisher, D., Eds.), pp. 529-587, Academic Press, New York. in