Plant Physiol. (1976) 58, 309-314
Isolation of Intact Chloroplasts and Other Cell Organelles from
Spinach Leaf Protoplasts1
Received for publication April 2, 1976 and in revised form May 21, 1976
MiKio NISHIMURA, DOUGLAS GRAHAM,2 AND TAKASHI AKAZAWA
Research Institute for Biochemical Regulation, School ofAgriculture, Nagoya University, Chikusa, Nagoya
464, Japan
ABSTRACT
Freshly prepared spinach leaf protoplasts were gently ruptured by
mechanical shearing followed by sucrose density gradient centrifugation
to separate constituent cell organelles. The isolation of intact Class I
chloroplasts (d = 1.21) in high yield, well separated from peroxisomes
and mitochondria, was evidenced by the specific localization of ribulose1,5-bisphosphate carboxylase (EC 4.1.1.39), NADP triose-P dehydrogenase (EC 1.2.1.9), and carbonic anhydrase (EC 4.2.1.1) in the fractions. A clear separation of chloroplastic ribosomes from the soluble
cytoplasmic ribosomes was also demonstrated by the band patterns of
constituent RNA species in the polyacrylamide gel electrophoresis. Localization of several enzyme activities specific to leaf peroxisomes, e.g.
catalase (EC 1.11.1.6), glycolate oxidase (EC 1.1.3.1), glyoxylate reductase (EC 1.1.1.26), glutamate glyoxylate aminotransferase (EC
2.6.1.4), seine glyoxylate aminotransferase, and alanine glyoxylate
aminotransferase (EC 2.6.1.12) in the peroxisomal fractions (d = 1.25),
was demonstrated. Overall results show the feasibility of the method for
the isolation of pure organelle components in leaf tissues.
sive trials, the isolation of pure chloroplast fractions retaining
both structural and functional integrity has reached only a limited success (31). A clear separation of pure leaf peroxisomes
free from other components has barely been accomplished (29).
Since cell wall materials are enzymically degraded during the
preparation of protoplasts, it is obvious that gentle breakage of
the latter surrounded only by the plasmalemma is ideal for
isolating the constituent organelles. In the work reported in this
communication we have undertaken the isolation of intact chloroplasts and other organelles from spinach leaf protoplasts in an
attempt to investigate the nature of the interorganellar relationship of photorespiration.
MATERIALS AND METHODS
Protoplasts. Protoplasts were prepared from freshly harvested
spinach leaves (Spinacia oleracea L var. Kyoho) essentially following the method reported previously (20). The only modification employed was that the concentration of mannitol was lowered to 0.7 M. The final preparations were shown to have
photosynthetic activities of 35 to 70 ,umol CO2 fixation/mg
Chl hr. The protoplasts (approximately 4 mg Chl) suspended in
2 ml of 0.05 M Tricine-NaOH buffer (pH 7.5) containing 0.5 M
sucrose and 0.1 % BSA were then ruptured through a syringe
A number of recent investigations have been increasingly (0.5 x 4 cm, Termo). A small piece of Miracloth (Calbiochem)
showing the potential usefulness of plant protoplasts for physio- was placed in the bottom of the needle (0.7 x 32 mm), and after
logical as well as biochemical research (25). In previous papers each stroke samples released from the needle were examined by
(20, 21) we have reported the value of spinach leaf protoplasts light microscope. Usually protoplasts were completely broken by
for studying the mechanism of photosynthesis and photorespira- three strokes (cf. Fig. 1).
Sucrose Density Gradient Centrifugation. The ruptured protion, the biggest advantage being the manipulation of homogeneous "naked leaf cell" preparations for such studies. We have toplast preparations (0.5 ml) were directly layered on top of 15
also emphasized that the maintenance of interorganellar rela- ml of the linear sucrose gradient (35-60%, w/w) dissolved in
tionships in protoplasts is useful for the analysis of the cellular 0.02 M Tricine-NaOH buffer (pH 7.5) and run at 24,000 rpm for
compartmentation mechanism in photorespiration which is be- 3 hr at 4 C using a Beckman-Spinco SW 25-3 rotor. At the end
lieved to proceed in multiorganellar fashion (28). As a next step, of the run, 0.5-ml fractions were collected and aliquots were
the separation of individual cell organelles having sufficiently used for measuring the enzymic activities as well as the content
high activities of enzyme reactions operating in photorespiration of Chl, protein, RNA, and DNA (see below).
and the subsequent reassembling are crucial to understanding
Enzyme Assays. (a) RuDP3 carboxylase (EC 4.1.1.39) activthe overall processes. However, both structural complexity and ity was measured as described by Nishimura et al. (22). (b)
fragility of organelles in leaf cells pose an obstacle for this task, NADP triose-P (EC 1.2.1.9) and NAD triose-P dehydrogenases
and breakage of structural organization of organelles during (EC 1.2.1.12) activities were determined by measuring the deisolation procedures is frequently encountered. Despite inten- crease in absorbance at 340 nm due to the oxidation of NADPH
or NADH by 3-PGA following the method of Heber et al. (10).
I This is
Paper No. XXXV in the series "Structure and Function of (c) Carbonic anhydrase (EC 4.2.1.1) activity was measured as
Chloroplast Proteins." The research is supported in part by the grants previously described by Atkins et al. (4) using veronal-pH indifrom the Ministry of Education of Japan, the Toray Science Foundation cator buffer at 0C and pH 8.2. (d) Catalase (EC 1.11.1.6)
(Tokyo), and the Naito Science Foundation (Tokyo).
activity was assayed by measuring the disappearance of H202
2Permanent address: Plant Physiology Unit, Commonwealth Scien- spectrophotometrically at 240 nm following the method of Luck
tific and Industrial Research Organization, Division of Food Research
and School of Biological Sciences, Macquarie University, North Ryde, (18). (e) Glycolate oxidase (EC 1.1.3.1) assay was carried out
N. S. W. 2113, Australia. Recipient of Award from the Japan Society
for the Promotion of Science under the Visiting Professorship Pro3Abbreviations: RuDP: ribulose 1,5-bisphosphate; DCPIP: 2,6-digramme (1974).
chlorophenolindophenol; 3-PGA: 3-P-glyceric acid.
309- Published by www.plantphysiol.org
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310
NISHIMURA, GRAHAM, AND AKAZAWA
anaerobically by measuring the reduction of DCPIP at 600 nm
following the method of Tolbert et al. (30). (f) NADH-glyoxylate reductase (EC1..1.1.26) activity was assayed by measuring
the decrease in absorbance at 340 nm due to the oxidation of
NADH by glyoxylate following the method of Zelitch and Gotto
(33). (g) Cyt c oxidase (EC 1.9.3.1) activity was determined by
measuring the decrease of reduced Cyt c at 550 nm according to
the method of Smith (26). (h) NAD-isocitrate dehydrogenase
(EC 1.1.1.41) activity was assayed by measuring the increase of
absorbance at 340 nm of NADH by isocitrate following the
method of Cox (6). (i) NAD-malate dehydrogenase (EC
1.1.1.37) activity was measured as described by Asahi and
Nishimura (3).(j) Glutamate glyoxylate aminotransferase (EC
2.6.1.4), serine glyoxylate aminotransferase, alanine glyoxylate
aminotransferase (EC 2.6.1.12), and aspartate a-ketoglutarate
aminotransferase (EC 2.6.1.1) activities were determined following the procedures described by Rehfeld and Tolbert (23).
Except in the case of carbonic anhydrase, enzyme assays were
carried out at 25 C for appropriate reaction periods, and activities are expressed as,mol substrates utilized or products
formed/minm tube (0.5 ml) (cf. Figs. 3 and 4).
Analytical Methods. Chl content was determined according to
the method of Arnon (2). Protein content was determined by the
spectrophotometric method of Lowry et al. (17) using BSA as a
standard.
The content of RNA and DNA in the fractionated samples
was determined according to the method of Fleck and Munro
(7).Each fraction (0.5 ml) was made to 5 ml with distilled H20
and 2.5 ml of cold 2.1 N HClI4 were added. After 15 min the
precipitate collected by centrifugation was washed twice with 0.1
ml of 0.7 N HC104. The precipitate was digested in an incubator
at 37C using 2 ml of 0.3 N KOH for 1 hr. At the end of
incubation, the samples were neutralized with 10 N HClO4 and
acidified with 1 volume of 1 N HClO4. The precipitate (DNA
fraction) separated by centrifugation was washed twice with 1 ml
of cold 0.5 N HCl04. The combined supernatant fluid and
washings (RNA fraction) were subjected to absorbance measurement at 260 nm and 275 nm. The RNA content was estimated by:
RNA (jig/ml) = 11.87 A26o nm -A275nm
The DNA fraction (precipitate) which was dissolved in 0.1 ml of
1 N KOH was assayed for DNA content by the diphenylamine
method.
Polyacrylamide Gel Electrophoresis. Each of the chloroplast
(No. 14 to 18 in Figure Sa) and the supernatant fractions (No.
27 to 31) were mixed with 1 volume of 90% phenol and stirred
vigorously for 15 min at room temperature. Afterward the
samples were thoroughly dialyzed against phenol-saturated water to remove sucrose. Then the samples were centrifuged and
the water phase fraction was carefully collected and 2 volumes of
ethanol were added. The precipitate formed was centrifuged and
washed twice with 4 ml of 70% ethanol. The precipitate was
dissolved in 100 ,ul of electrophoretic buffer and applied to the
polyacrylamide gel electrophoresis following the method of
Loening and Ingle (16). The whole leaf RNA fraction was
extracted from spinach leaves by 50% phenol and treated in the
same way. After electrophoresis (5 mamp/tube, 2 hr), the gel
was stained with methylene blue. A Joyce Loeble Chromo-Scan
was used to obtain the densitometric tracings of the gels stained
with dye.
RESULTS
Isolation of Intact Chloroplasts. The primary purpose of the
present investigation was to isolate preparations of intact chloroplast after breakage of protoplasts. Several procedures for
breakage were examined such as osmotic shock and mechanical
Plant
Physiol. Vol. 58,
1976
a syringe.
breakage by Teflon homogenizer or passage through
The gentle mechanical shearing using a syringe was found to be
most satisfactory. Figure 1 shows the release of a homogeneous
population of chloroplasts examined by light microscope after
the syringe treatment. A single passage is compared with a triple
passage preparation. By phase contrast microscopy thethechloroouter
with a halo around
plasts appeared brightly refractive,
(27). A clear separation of a single band
margin of the particles
of chloroplasts (d = 1.21), practically free from stripped chloroplasts, was achieved by sucrose density gradient centrifugation
(Fig. 2), a nearly complete recovery of chloroplast fractions was
achieved by this procedure (93 % recovery from starting proto-
plasts on Chi basis).
Cellular Localization of Enzyme Activities in Chloroplasts and
Other Organelles. A satisfactory resolution of chloroplasts and
other constituent organelles can be supported from experimental
results examining the enzyme localization in separated fractions.
Figure 3 shows a sedimentation profile of the broken protoplasts
The
preparations after sucrose density gradient centrifugation. coinmarker enzyme exactly
prominent peaks of chloroplastic
with the peak of
(d = 1.21) (Fig. 3, a and b). Some
activities of RuDP carboxylase and NADP triose-P dehydrogenase are detectable in the soluble supernatant fraction, but they
activities of
comprise only a small proportion of the total. Sizable
carbonic anhydrase were found to be clearly associated with
top, soluble
chloroplasts, but much activity remained in thefraction
reprefractions. Presumably enzyme activities in each
sent the chloroplastic and cytoplasmic carbonic anhydrase (8),
but that in the soluble fraction is so high that contamination by
enzyme leached out, or from the surface, of chloroplasts must be
Chl
cide
considered.
Marker
enzyme
activities of peroxisomes
(28,
30, 32), i.e.
catalase, glycolate oxidase, and glyoxylate reductase, are clearly
present in peroxisome fractions (d = 1.25), although much of
these activities (approximately 70%) are also present in the
due to the
supernatant fractions (Fig. 3c). This is probably
and leaching of the
fragility of the single membranous organelles
enzyme during the separation procedure. Tolbert reported (29)
that leaf peroxisomes comprise 1 to 1.5% of the total protein,
and the present results show that the content of peroxisomes
recovered is approximately 3% on a protein basis (Fig. 3a).
the osmotic treatment of protoplasts followed by
Employing
sucrose density gradient centrifugation, both glycolate oxidase
and glyoxylate reductase activities were almost exclusively located in the top, soluble fractions of the centrifuged samples,
although chloroplasts and mitochondria were fairly well preserved.
While not much data are available regarding the separation of
mitochondria from other organelles in leaf tissues, reported
results from several laboratories are somewhat conflicting (11,
23, 24, 32). We have consistently observed that when the protowithout BSA
plast preparations suspended in sucrose solution
were subjected to mechanical breakage followed by gradient
fractions (d = 1 .22) overcentrifugation, the mitochondrial
(d = 1.21), probably because of theof
lapped withofchloroplasts
absorption solute (sucrose) through the outer membranes
mitochondria (T. Asahi, personal communication). Activity of
chloroCyt c oxidase was found in the leading edge fractionsbeofbasically
results are judged to
plasts (data not shown). These
similar to those of Rocha and Ting (24) and Huang and Beevers
(1 1); in such studies the density of mitochondria (d = 1.21) was
shown to be larger than that of stripped chloroplasts (d = 1 .17).
On the other hand, we have found that inclusion of BSA in the
mitochondria,
suspending media exhibits a preservative effect on
the density of mitochondria becoming distinctly smaller
thereby
than that of the chloroplasts. Figure 3d shows the specific localization of both Cyt c oxidase and NAD-isocitrate dehydrogenase
in the fractions of d = 1.18. Figure 3e shows that activities of
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Copyright © 1976 American Society of Plant Biologists. All rights reserved.
Plant
Physiol. Vol. 58,
~
311
ORGANELLE SEPARATION FROM PROTOPLASTS
1976
©
1 Stroke
1s
*
N@
,t
_
--
/.-,w+w.?1
I4.
i
ct.* 'o
.*. 0. -:,-
-s
.
V
s
+8
P
.w
.,,jl,
,
,.
(x 5OO)
®i 3 Strokes
j
#-t
r
I
.
P
'
t
r.
Ob
9
f) 5''
r.
rs
X,
-ifs
.0
II"
a&
Jsz
ekAf&
8'.
1.
4
&.o
n
tr~*
frt
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1.
.9
S
4-J
/ 'I
.r
p~~
(
e-
0r
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(x
'
.
1000)
FIG. 1. Photomicrographs of gently ruptured protoplasts. a and b show the spinach protoplasts ruptured by gentle shearing through syringe
and three times, respectively. In a arrows indicate unbroken protoplasts. Experimental procedures are described in the text.
NAD-malate dehydrogenase are detected in peroxisomes, mitochondria, and chloroplasts. In agreement with results of previous
investigators (28, 32), the specific activity is highest in peroxisomes, but it must be pointed out that the enzyme activity is
clearly detected in the chloroplastic fractions.
Activities of four aminotransferases which are reported to be
operative in the glycolate pathway were assayed in the separated
fractions and results are shown in Figure 4. In accordance with
the results of Tolbert and his associates (23, 28, 32), a tight
association of three aminotransferases, i.e. glutamate glyoxylate
(Fig. 4a), serine glyoxylate (Fig. 4b), and alanine glyoxylate
aminotransferases (Fig. 4c), with peroxisomes was clearly demonstrated. However, for each of these enzymes some activity was
once
detected in the soluble supernatant fractions and small activities
in other particulate fractions as well. Activity of aspartate aketoglutarate aminotransferase was detected in all three organelles: chloroplasts, peroxisomes, and mitochondria (Fig. 4d),
presumably representing each individual isozyme (23, 32).
Isolation of Ribosomal RNA. An additional proof for the
isolation of intact chloroplasts can be seen from the separation of
chloroplastic and cytoplasmic ribosomal RNA. The profile of
RNA analysis in the separated fractions is given in Figure 5a. In
order to characterize the RNA components of the chloroplast
and soluble cytoplasmic fractions, RNA sepcies were resolved by
polyacrylamide gel electrophoresis according to the method of
Loening and Ingle (16). The stained electrophoresed gel samples
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Copyright © 1976 American Society of Plant Biologists. All rights reserved.
312
NISHIMURA, GRAHAM, AND AKAZAWA
Plant
Physiol. Vol. 58,
O
1976
trifugation of gently homogenized leaf cells using isotonic sorbitol solution (12, 31). It is inevitable that all such preparations
are mixed populations of various organelles. It is unavoidable
that these methods cause some structural damage of chloroplasts
and other organelles during the step of breaking the leaf tissues.
The use of leaf protoplasts as a starting material is different in
principle from the previous commonly used methods, and the
present investigation clearly shows that the gentle breakage of
naked leaf cells (protoplasts) followed by sucrose density centrifugation is advantageous for separating three major organelles,
namely chloroplasts, mitochondria, and peroxisomes, in pure
form. The specific localization of some enzyme activities as well
as the ribosomal RNA in the distinctly separated chloroplast
hi
oroplasts
3
1
a 2
8
I
r
FIG. 2. Photograph of the separation of chloroplasts by sucrose density gradient centrifugation. Mechanically broken protoplasts (Fig. lb)
(approximately 2 mg Chl) were applic to the sucrose density gradient
centrifugation as explained in the text
and their scan tracing are given in Figure 6 (b and c), in comparison with the picture of the total RNA fraction obtained from the
whole spinach leaf (Fig. 6a). The pictures clearly show that the
chloroplastic RNA (23s, 16s) is well separated from the cytoplasmic RNA (25S, 18s), indicating that the chloroplastic and
cytoplasmic fractions contain chloroplastic rRNA and cytoplasmic rRNA, respectively. Percentage distribution of each RNA
species in the individual fraction was calculated from the figure
and the values are summarized in Table I. A small amount of
cross-contamination between the two fractions can be seen, but
we cannot exclude a possibility that cytoplasmic rRNA is associated with the chloroplastic outer membranes (1). The same type
of association is clearly demonstrated in yeast mitochondria
(13). Also, the presence of small amounts of unknown RNA
species having smaller molecular size can be seen in Figure 6, b
and c, and is probably due to the degradation of each respective
rRNA species during the preparation of samples (16).
Figure Sb shows the results of DNA analysis. There are two
major DNA-containing fractions, I and II, but none of them is
fully characterized. Although we found that fraction I is positive
to the Feulgen staining, the sedimented pellets in centrifuge
tubes are also positive to this stain. It will be noted that since the
absorption spectra of peak II fractions analyzed by the diphenylamine method showed the presence of two prominent peaks at
650 nm and 600 nm, there is a possibility that non-DNA substances are present in these fractions.
DISCUSSION
The hitherto reported structurally intact and functionally active chloroplasts were preparations obtained by low speed cen-
-
8
-1
il
(botrom) 5
10
15
FRACTION
20 25
NUMBER
0
0
o
o
0
5
30 (top)
FIG. 3. Localization of enzyme activities in the separated fractions
after sucrose density gradient centrifugation of mechanically ruptured
protoplast preparations. Mechanically ruptured protoplasts were subjected to sucrose density gradient centrifugation as shown in Fig. 2, and
the 0.5-ml fractions separated were used for the following assays: a:
protein and Chl; b: RuDP carboxylase, NADP triose-P dehydrogenase,
and carbonic anhydrase; c: catalase, glycolate oxidase, and glyoxylate
reductase; d: Cyt c oxidase and NAD-isocitrate dehydrogenase; and e:
NAD-malate dehydrogenase and NAD triose-P dehydrogenase. Experimental details for the enzyme activity measurements are described in the
text. Except in the case of carbonic anhydrase all other enzyme activities
m tube
are expressed as ,umol substrates utilized or products formed/min
(0.5 ml).
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Plant
Physiol. Vol. 58,
313
ORGANELLE SEPARATION FROM PROTOPLASTS
1976
*12 Al]
I
8
f4 *
deterioration of their biochemical activities. The light-dependent
CO2 fixation activity is usually low. The final chloroplast preparations (d = 1.21) obtained in the present study exhibited
fixation of about 5 Mumol C02/mg Chl hr. Therefore, in spite of
the fact that chloroplasts appear to retain their structural integrity microscopically, the type of preparations isolated can be
classified as Class I according to Spencer and Unt (27). An
additional factor causing the decline of photosynthetic activity is
likely to be the time lapse during preparation of protoplasts (4
hr), probably resulting in the consumption of photosynthetic
intermediates during this step. It will be recalled that previously
-
25s
®Whole leaf
/
4
(boo) 5
j2
0
1
2
0
0(t
18s
A--
I1 hQ-.YkoOY/
fRACTION
16s
NUMBER
FIG. 4. Localization of various aminotransferase activities in the separated fractions after sucrose density gradient centrifugation of mechanically ruptured protoplast preparations. The preparation used for mechanical breakage and sucrose density gradient centrifugation was the
same as that shown in Fig. 3. Aliquots of separated fractions were
subjected to the following enzyme assays: a: glutamate glyoxylate aminotransferase; b: serine glyoxylate aminotransferase; c: alanine glyoxylate aminotransferase; and d: aspartate a-ketoglutarate aminotransferase. Experimental details for the enzyme activity measurements are
described in the text, and all enzyme activities are expressed as ,umol
products formed/min- tube (0.5 ml).
I
(ID Fraction 14-18 23s
13s
16s
© Fraction 27-31
25s
8s
(bonom) 5
10
15
FRACTION
20
25
30 (top)
NUMBER
FIG. 5. Localization of RNA and DNA in the separated fractions
after sucrose density gradient centrifugation of mechanically ruptured
protoplast preparations. The preparation used for mechanical breakage
and sucrose density gradient centrifugation was the same as that of Fig.
2. Aliquots of separated fractions were analyzed for (a) RNA and (b)
DNA.
fractions (d = 1.21) is evidence that the isolated fractions are
pure. However, the present method has its own limitations. As is
generally recognized, gradient separation of chloroplasts on a
dense sucrose solution for a prolonged period (3 hr) causes
FIG. 6. Polyacrylamide gel electrophoresis of RNA species. (a)
Whole leaf RNA, (b) chloroplastic RNA obtained from pooled fraction
(No. 14 to 18 of Fig. Sa), and (c) soluble cytoplasmic RNA obtained
from pooled fractions No. 27 to 31 of Fig. 5a were subjected to polyacrylamide gel electrophoresis according to the method of Loening and
Ingle (16). After separation of the constituent RNA species, the gels
were stained with methylene blue and the density tracing was made.
Experimental details are described in the text.
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NISHIMURA, GRAHAIM, AND AKAZAWA
314
Table I. Percentage Distribution of RNA Species
Values are computed from the scanned figures shown in Figure 6.
Whole leaf
Chloroplastic fraction
(tubes 14-18)
cytoplasmic
Soluble
fraction (tubes 2731)
25s
23s
18s
16s
Species
36.2
5.6
18.5
36.4
25.7
3.2
13.5
28.0
6.1
26.8
35.2
2.8
22.0
3.3
26.5
several investigators attempted to enhance the CO2 fixation
capacity of chloroplast preparations by supplementing some P
esters, e.g. fructose-diP, ribose-5-P, and 3-PGA (5). Furthermore, one can assume that the contact of cytoplasmic and other
cellular inclusions with chloroplasts during the preparative step
of chloroplasts may result in both enzymic and nonenzymic
degeneration of the membranous structure of the latter. Recently reported procedures for the isolation of active chloroplast
preparations are all based on very rapid separation using a
medium such as silica gel (19) or two phase dextran-polyethylene glycol (14), but the homogeneity of separated organelle
fractions has not been well documented in these studies. Although the present procedure appears to be far superior to
others to the extent of isolating enzymically and structurally pure
chloroplasts, further technical improvements are needed in order to obtain fully functional Class A chloroplasts (9).
It is known that single membranous organelles such as peroxisomes are particularly fragile and are difficult to isolate in the
intact form. Lips (15) reported that enzyme content of plant
microbodies is liable to be affected by experimental procedures
such as homogenization or sedimentation. It must be pointed out
that there are no satisfactory criteria for the intactness of peroxisomes. The present method is considered to be useful for separating them free from other cellular constituents. From the
specific localization of seven major component enzymes with the
fractions of d = 1.25 the separation of intact peroxisomes free
from other organelles can be achieved by the present technique.
However, some enzymic activities are leached out into the soluble fractions located at the top of the gradient, whereas they may
represent specific location in the cytosol in vivo.
Note. After completion of this manuscript our attention was
drawn to the paper by Rathnam and Edwards which described
the isolation of chloroplasts from various grasses (Rathnam, C.
K. M. and G. E. Edwards. 1976. Protoplasts as a tool for
isolating functional chloroplasts from leaves. Plant Cell Physiol.,
17: 177-186).
Acknowledgments -We wish to record our hearty thanks to H. Beevers for his invaluable
discussion and advice during the course of this investigation and to A. Watanabe for the guidance
of polyacrylamide gel electrophoresis technique.
LITERATURE CITED
1. APEL, K. AND H. G. SCHWEIGER. 1973. Site of synthesis of chloroplast membrane proteins.
Evidence for three types of ribosomes engaged in chloroplast protein synthesis. Eur. J.
Biochem. 38: 373-383.
Plant Physiol. Vol. 58, 1976
2. ARNON, D. I. 1949. Copper enzymes in chloroplasts. Polyphenol oxidase in Beta vtlgaris.
Plant Physiol. 24: 1-15.
3. ASAHI, T. AND M. NISHIMURA. 1973. Regulatory function of malate dehydrogenase
isoenzymes in the cotyledons of mung bean. J. Biochem. 73: 217-225.
4. ATKINS, C. A., B. D. PAT-rERSON, AND D. GRAHAM. 1972. Plant carbonic anhydrases. 1.
Distribution of types among species. Plant Physiol. 50: 214-217.
5. BAMBERGER, E. S. AND M. GIBBs. 1965. Effect of phosphorylated compounds and inhibitors on CO, fixation by intact spinach chloroplasts. Plant Physiol. 40: 919-926.
6. Cox. G. F. 1969. Isocitrate dehydrogenase (NAD-specific) from pea mitochondria. Methods Enzymol. 13: 47-51.
7. FLECK, A. AND H. N. MUNRO. 1962. The precision of ultraviolet absorption measurements
in the Schmid-Thannhauser procedure for nucleic acid estimation. Biochim. Biophys. Acta
55: 571-583.
8. GRAHAM, D., G. A. PENY, AND C. A. ATKINS. 1974. In search of a role for carbonic
anhydrase in photosynthesis. In: R. L. Bieleski, A. R. Ferguson, and M. M. Gresswell,
eds., Mechanisms of Regulation of Plant Growth. Bulletin 12. The Royal Society of New
Zealand, Wellington. pp. 251-258.
9. HALL, D. 0. 1972. Nomenclature for isolated chloroplasts. Nature New Biol. 235: 125126.
10. HEBER, U., N. G. PON. AND M. HEBER. 1963. Localization of carboxydismutase and triose
phosphate dehydrogenase in chloroplasts. Plant Physiol. 38: 355-360.
11. HUANG, A. H. C. AND H. BEEVERS. 1971. Isolation of microbodies from plant tissues. Plant
Physiol. 48: 637-641.
12. JENSEN, R. G. AND J. A. BASSHAM. 1966. Photosynthesis by isolated chloroplasts. Proc.
Nat. Acad. Sci. U. S. A. 56: 1095-1 101.
13. KELLEMS, R. E., U. F. ALLISON. AND R. A. BUTOW. 1974. Cytoplasmic type 80 s
ribosomes associated with yeast mitochondria. II. Evidence for the association of cytoplasmic ribosomes with the outer mitochondrial membrane in situ. J. Biol. Chem. 249: 32973303.
14. LARSSON, C. AND P. A. ALBERTSSON. 1974. Photosynthetic "4CO2 fixation by chloroplast
population isolated by a polymer two-phase technique. Biochim. Biophys. Acta 357: 412419.
15. LipS, S. H. 1975. Enzyme content of plant microbodies as affected by experimental
procedures. Plant Physiol. 55: 598-601.
16. LOENING, U. E. AND J. INGLE. 1967. Diversity of RNA components in green plant tissues.
Nature 215: 363-367.
17. LOWRY, 0. H., N. J. ROSEBROUGH, A. L. FARR, AND R. J. RANDALL. 1951. Protein
measurement with Folin phenol reagent. J. Biol. Chem. 193: 265-276.
18. LUCK, H. 1965. Catalases. In: H. Y. Bergmeyer. ed., Methods of Enzymatic Analysis.
Academic Press, New York. pp. 885-894.
19. MORGENTHALER, J. J., C. A. PRICE, J. M. ROBINSON, AND M. GIBBS. 1974. Photosynthetic
activity of spinach chloroplasts after isopycnic centrifugation in gradients of silica. Plant
Physiol. 54: 532-534.
20. NISHIMURA, M. AND T. AKAZAWA. 1975. Photosynthetic activities of spinach leaf protoplasts. Plant Physiol. 55: 712-716.
21. NISHIMURA, M., D. GRAHAM, AND T. AKAZAWA. 1975. Effect of oxygen on photosynthesis
by spinach leaf protoplasts. Plant Physiol. 56: 718-722.
22. NISHIMURA, M., T. TAKABE, T. SUGOYAMA, AND T. AKAZAWA. 1973. Structure and
function of chloroplast proteins. XIX. Dissociation of spinach leaf ribulose-1.5-diphosphate carboxylase by p-mercuribenzoate. J. Biochem. 74: 945-954.
23. REHFELD, D. W. AND N. E. TOLBERT. 1972. Aminotransferases in peroxisomes from
spinach leaves. J. Biol. Chem. 247: 4803-4811.
24. ROCHA, V. AND I. P. TING. 1970. Preparation of cellular plant organelles from spinach
leaves. Arch. Biochem. Biophys. 140: 398-407.
25. RUESINK, A. W. 1971. Protoplasts of plant cells. Methods Enzymol. 23: 197-209.
26. SMITH, Z. 1955. Spectrophotometric assay of cytochrome c oxidase. Methods Biochem.
Anal. 2: 427-434.
27. SPENCER, D. AND H. UNT. 1965. Biochemical and structure correlations in isolated spinach
chloroplasts under isotonic and hypotonic conditions. Aust. J. Biol. Sci. 18: 197-2 10.
28. TOLBERT, N. E. 1971. Microbodies-peroxisomes and glyoxysomes. Annu. Rev. Plant
Physiol. 22: 45-74.
29. TOLBERT, N. E. 1971. Isolation of leaf peroxisomes. Methods Enzymol. 23: 665-682.
30. TOLBERT, N. E., A. OESER, T. KISAKI, R. H. HAGEMAN, AND R. K. YAMAZAKI. 1968.
Peroxisomes from spinach leaves containing enzymes related to glycolate metabolism. J.
Biol. Chem. 243: 5179-5184.
31. WALKER, D. A. 1971. Chloroplasts (and Grana): aqueous (including high carbon fixation
ability). Methods Enzymol. 23: 211-220.
32. YAMAZAKI, R. K. AND N. E. TOLBERT. 1970. Enzymatic characterization of leaf peroxisomes. J. Biol. Chem. 245: 5137-5144.
33. ZELrrCH, I. AND A. M. GoTTo. 1962. Properties of a new glyoxylate reductase from leaves.
Biochem. J. 84: 541-546.
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