J. Cell Sci. 3, 327-340 (1968)
Printed in Great Britain
327
AUTORADIOGRAPHIC EVIDENCE FOR THE
IN SITU SYNTHESIS OF CHLOROPLAST
AND MITOCHONDRIAL RNA
SARAH P. GIBBS*
M.R.C. Epigenetics Research Group,
Institute of Animal Genetics, Edinburgh
SUMMARY
The rate of appearance of labelled RNA in the chloroplast and mitochondria as compared
with the rate in the remaining cytoplasm was studied in the unicellularflagellate,Ochromonas
danica, by electron-microscope autoradiography. Greening cells were labelled withuridine5,6['H] for a short (30 min) and a long (2 h) interval and the concentration of label, expressed as
grains/unit area, determined for each cell component. The data demonstrate that there is the
expected lag in the labelling of the cytoplasm proper, but no apparent lag in the labelling of the
chloroplast and mitochondria. This observation, combined with the fact that after the short
labelling time the chloroplast and mitochondria have a much heavier concentration of labelled
RNA than the surrounding cytoplasm, indicates that most, if not all, chloroplast and mitochondrial RNA is synthesized m situ.
The three kinds of ribosomes present in the cell are distinctly different in size. The mitochondrial ribosomes measure 150-170 A in diameter, the chloroplast ribosomes average
170-200 A in diameter, whereas the cytoplasmic ribosomes are 210-230 A in diameter in
glutaraldehyde-osmium-fixed cells. During chloroplast development in the light, the number
of chloroplast ribosomes increases approximately tenfold.
INTRODUCTION
During the past 5 years it has become well established that two cytoplasmic
organelles, mitochondria and chloroplasts, contain their own species of DNA which
differ in a number of respects from nuclear DNA. The presence of DNA in mitochondria was first indisputably demonstrated by Nass & Nass (1963 a, b) using enzyme
digestion studies combined with electron microscopy. The following year Luck &
Reich (1964) succeeded in isolating mitochondrial DNA from Neurospora, and subsequently mitochondrial DNA has been isolated from a variety of organisms (see
Granick & Gibor, 1967). Ris & Plaut (1962) were the first to demonstrate that chloroplasts contain fine DNase-digestible fibrils. The next year Chun, Vaughan & Rich
(1963), using equilibrium density-gradient centrifugation, showed that DNA isolated
from chloroplasts had a different buoyant density than nuclear DNA. Since then,
chloroplast DNA has been demonstrated in a wide variety of plants (see Iwamura,
1966).
• Present address: Electron Microscope Unit, Stewart Biology Building, McGill University,
Montreal, Canada.
328
S. P. Gibbs
There is also good evidence that both chloroplasts and mitochondria contain ribosomal and transfer RNA and indirect evidence that they contain messenger RNA.
Ribosomes have been isolated from the chloroplasts of a number of species of plants
and have been found to have a lower sedimentation coefficient than cytoplasmic
ribosomes, 70s instead of 80s (Lyttleton, 1962; Clark, Mathews & Ralph, 1964;
Sissakian, Filippovich, Svetailo & Aliyev, 1965; Spencer, 1965; Boardman, Francki &
Wildman, 1966). In addition, Jacobson, Swift & Bogorad (1963) have demonstrated
that the dense 170 A granules of the chloroplast matrix are digestible with ribonuclease. Small RNase-digestible granules have also been observed recently in mitochondria by several investigators (Swift, Adams & Larsen, 1964; Andre & Marinozzi,
1965; Schuster, 1965; Leduc, Bernhard & Tournier, 1966). These mitochondrial
ribosomes have proved more difficult to isolate free from contamination by cytoplasmic
ribosomes than are chloroplast ribosomes, but recently O'Brien & Kalf (1967 a, b)
have obtained preparations of 55 s ribosomes from rat liver mitochondria. Isolated
chloroplasts have been shown to contain a 4 s RNA (Spencer & Whitfeld, 1966;
Berger, 1967) which has acceptor activity for amino acids (Sissakian et al. 1965). In
the case of mitochondria, Barnett & Brown (1967) have recently demonstrated that
mitochondria isolated from Neurospora contain amino acid-acceptor transfer RNA's
for 19 amino acids and that in at least one case, that of aspartic acid transfer RNA, the
mitochondria contain a unique species of this transfer RNA which is absent from the
cytoplasm.
The presence of messenger RNA in higher plant plastids can be inferred from the
fact that polyribosomes are often present (Clark, 1964; Gunning, 1965; Newcomb,
1967; Stutz & Noll, 1967) and that they are more active in incorporating amino acids
into proteins than are the 70 s ribosomes (Chen & Wildman, 1967). Brawerman &
Eisenstadt (1964) have also presented evidence for the presence of messenger RNA in
Euglena chloroplasts. As yet messenger RNA in mitochondria has not been directly
demonstrated, but its presence can be inferred from the fact that isolated mitochondria are capable of incorporating amino acids into proteins (see Wintersberger,
1965)One of the major questions which arises from the observation that chloroplasts and
mitochondria contain nucleic acids is, 'Are the various species of chloroplast and
mitochondrial RNA synthesized in situ using chloroplast and mitochondrial DNA,
respectively, as primer, or are they synthesized in the nucleus, coded by nuclear
DNA?' There is now a substantial amount of biochemical evidence that at least some
of the RNA present in chloroplasts and mitochondria is coded in situ. Actinomycin D
inhibits RNA synthesis to varying degrees in both chloroplast (Kirk, 1964; Schweiger
& Berger, 1964; Shah & Lyman, 1966) and mitochondrial preparations (Luck &
Reich, 1964; Kalf, 1964; Neubert & Helge, 1965; Wintersberger & Tuppy, 1965;
Wintersberger, 1966). In addition, in vivo studies on Acetabularia have shown that
newly synthesized RNA appears in the chloroplasts of enucleated plants (Shephard,
1965; Janowski, 1965). Schweiger, Dillard, Gibor & Berger (1967) and Berger (1967),
also using enucleated cells of Acetabularia, have shown that both chloroplast ribosomal
RNA and chloroplast transfer RNA are synthesized within the chloroplast. Very
RNA synthesis in chloroplasts and mitochondria
329
recently, Scott & Smillie (1967) have shown by DNA-RNA hybridization techniques
that the chloroplast ribosomes of Euglena are coded by chloroplast DNA.
In the present study, the question of the site of synthesis of chloroplast and mitochondrial RNA has been investigated by means of autoradiography. A number of
earlier autoradiographic studies showed that when cells are fed radioactive RNA precursors, the initial labelling appears in the nucleus and only after a time lag does any
appreciable radioactivity appear in the cytoplasm (see Prescott, 1964, for a review).
These studies were generally interpreted to mean that the synthesis of all RNA in the
cell takes place within the nucleus. However, in light of the rapidly accumulating
biochemical evidence that mitochondrial and chloroplast RNA is synthesized in situ,
it is necessary to repeat these experiments to determine whether there is also a lag in
the incorporation of labelled RNA precursors into the RNA of mitochondria and
chloroplasts or whether these organelles label rapidly from the beginning as one
would expect if they synthesized their own RNA independently of the nucleus.
There are two important considerations in planning such a study. First it is necessary to employ the techniques of quantitative electron-microscopic autoradiography
in order to be able to distinguish mitochondrial from other cytoplasmic labelling with
certainty. Secondly, in view of the fact that chloroplast and mitochondrial RNA often
form only a small fraction of the total cellular RNA, it is desirable to pick a system in
which a cell is preferentially synthesizing chloroplast and mitochondrial RNA. This
can easily be done in the case of chloroplasts by choosing a system in which chloroplast
growth and development is induced by light. It is more difficult to find a system in
which a cell is also preferentially synthesizing mitochondrial RNA, but one can at
least be certain that mitochondria are synthesizing (or receiving) some RNA if dividing
cells are employed. Consequently, for these experiments log phase cells of the flagellate Ochromonas danica (division time: 14 h) were used at two different times during
the period of rapid chloroplast development. These cells were exposed to tritiated
uridine for a short and a long period and the amount of labelled RNA in each cell
organelle was determined using electron-microscopic autoradiography.
MATERIALS AND METHODS
Biological material
Stocks of Ochromonas danica Pringsheim (1955) were obtained from the Culture Collection
of Algae and Protozoa at Cambridge. Cells were grown at 27 °C in defined medium (Aaronson
& Baker, 1959) either in light-tight boxes or under incandescent lamps at an intensity of 200 ft-c.
Cell counts and chlorophyll analyses were made as described previously (Gibbs, 1962).
Electron microscopy
For general observations, cells from a variety of dark-grown and greening cultures were fixed
for 3 h in cold 2-5% glutaraldehyde in O-IM phosphate buffer, pH 7-2. Following 3 30-min
rinses in buffer alone, the cells were post-fixed for 2 h in cold 2 % osmium tetroxide, also in
o-i M phosphate buffer, rinsed twice in buffer, and embedded in 2 % Nobel's agar. Small agar
blocks were cut, dehydrated in an ethanol series, followed by epoxy-propane, and embedded
in Araldite. Sections were stained for 30 min with lead citrate (Reynolds, 1963) and viewed
with an AEI EM 6 electron microscope.
330
S. P. Gibbs
Autoradiography
For the uridine-labelling experiments, cultures of Ochromonas were grown in the dark 6 or 7
days and then placed in the light to green, either for 2 h (culture A) or for 24 h (culture B)
prior to the addition of the isotope. Previous studies (Gibbs, 1962) had shown that cultures in
the logarithmic phase of growth synthesized chlorophyll much more rapidly than those in the
linear or stationary phase. Consequently dark-grown cultures were selected for these experiments which were near the end of the log phase of growth and contained approximately 8x10'
cells per ml.
Following the pre-illumination period, each culture was concentrated 20-fold by centrifuging
at 2000 rev/min for 3 min and resuspending the cells in fresh medium. Microscopic examination
ascertained that no cells were broken or rendered immobile by ccntrifugation. One-ml aliquots
of concentrated cells were pipetted into 5 ml Erlenmeyer flasks and 50 fie of uridine-5,6['H]
dissolved in 0-2 ml distilled water added to each aliquot (isotope from the Radiochemical
Centre, Amersham; specific activity, 3 c/mmole). The flasks of cells plus isotope were gently
shaken in the light for 30 min and 2 h respectively. The incorporation was terminated by
flooding the cells with a large excess offixative(1 % osmium tetroxide in acetate-veronal buffer,
pH 7-4). The cells were fixed for 7 h at 2 °C, rinsed thoroughly in 3 changes of buffer, blocked
in agar, dehydrated in ethanol, and embedded in prepolymerized methacrylate.
For autoradiography uniform silver sections were cut and picked up on colloidion-coated
nickel grids. The grids were attached to glass slides and coated with a monolayer of recently
manufactured Ilford L-4 emulsion, using the loop method of Caro & van Tubergen (1962).
The slides were stored at 2 °C in Bakelite slide boxes containing a small cylinder of Drierite.
After 5J months exposure, the slides were developed in D-19 for 5 min at 17-5 °C.
Cells from the 24-h-light culture (culture B) which had been labelled with uridine-5,6[5H]
for 30 min and for 2 h were used for the ribonuclease digestion studies. After fixation in i-6 %
glutaraldehyde in o-i M phosphate buffer, pH 7-2, for 30 min, the cells were blocked in agar and
embedded in glycol methacrylate by the method of Leduc, Marinozzi & Bernhard (1963).
Polymerization was carried out at 15 °C using an u.v. lamp. Thin sections were cut, mounted on
colloidion-coated grids, and prepared for autoradiography after:
(a) Digestion with ribonuclease A (Sigma, 5 x crystallized), 1 mg/ml, pH 6-7, for 30 min
at 38 °C.
(b) Digestion in ribonuclease as above, followed by a 1 min rinse in cold 5 % TCA.
(c) Digestion in distilled water, adjusted to pH 6'7, for 30 min at 38 CC.
(d) Digestion in distilled water as above, followed by a 1 min rinse in cold 5 % TCA.
(«) No treatment.
These grids were exposed for 6J months and developed with D-19 as above.
For comparison purposes, a single experiment was done in which the cells were labelled
with thymidine-[aH]methyl (the Radiochemical Centre, specific activity, 14-9 c/mmole). In
this experiment, dark-grown, log phase cells were pre-illuminated for 5-5 h. Then 50 fie
[*H]thymidine dissolved ino-i ml distilled water was added to I ml of concentrated cells, and
the cells exposed to the light in the presence of isotope for a further 18 h. Fixation and embedding of these cells was by the glutaraldehyde—osmium—Araldite method detailed above. Radioautographs were prepared by the same methods used in the uridine experiments, except that a
longer exposure time (6J months) was required.
Quantitation of results
In order to quantitate the results, it was necessary to take micrographs at random. This is
difficult to do as there is a strong temptation to photograph only well-fixed and well-labelled
cells. On each grid, therefore, an area was selected where the sections were uniformly thin, and
every cell or field of cells in a section was photographed until approximately 20 micrographs
were obtained. If 20 micrographs were not obtained from a single section, another section
farther down the ribbon was similarly surveyed. After 20-25 exposures, a new grid was selected.
Grids were photographed in this manner until at least 100 good micrographs were obtained for
each of the 4 variables of the experiment. All micrographs were taken at an initial magnification
of x 6000 and were printed on 8 x 10 in. paper at a final magnification of x 16000. In counting
RNA synthesis in chloroplasts and mitochondria
331
grains, an arbitrary rule was made that if over half of a grain fell within the boundary of an
organelle, then the grain originated in that organelle. Grains were counted on the 8 x 10 in.
prints, but it was frequently necessary to check the grain counts on the negative, since a silver
grain can sometimes be confused with dirt on the print, but never on the negative.
In order to determine the relative volume occupied by each cell organelle, each micrograph
was traced in its entirety on high-quality tracing paper, and each organelle was carefully cut out,
collected in groups, and weighed. A series of micrographs, such as those used, taken at random
of a uniform population of cells is equivalent to serial sections through a single representative
cell. (Actually for each of the 4 variables the equivalent of serial sections through 2 entire cells
were traced.) That relative volume may be thus estimated from measurements of area was early
observed by the French geologist, Delesse, and has recently been demonstrated mathematically
by Chayes (1965).
RESULTS
Electron-microscopic observations
Chloroplast ribosomes. The matrix of the chloroplast of greening cells of Ochromonas
contains numerous ribosomes, which average 170-200 A in diameter in glutaraldehydeosmium fixed cells (Fig. 1). The cytoplasmic ribosomes are distinctly larger (210230 A) than the chloroplast ribosomes. The chloroplast ribosomes are most abundant
in the areas of matrix lying between the three-disc lamellar bands, and are relatively
sparse in the narrow layer of matrix lying between the outermost lamellar band and the
chloroplast envelope (Fig. 1). The majority of the chloroplast ribosomes lie free in the
chloroplast matrix; some, however, appear to be attached to the outer membranes of
the three-disc bands.
An important question in considering what kinds of RNA are synthesized in developing chloroplasts is whether or not there is an increase in the number of chloroplast ribosomes during chloroplast development. Figure 2 illustrates a typical proplastid of a dark-grown cell. The matrix contains scattered ribosomes of the same size
as those in green chloroplasts. Since in green chloroplasts all the chloroplast ribosomes
are crowded together in the areas of matrix remaining between the lamellar bands, it is
difficult to estimate by eye whether or not there is an increase in the number of
ribosomes per area during chloroplast development. Counts made on randomly
selected i-/t2 areas showed that the proplastids of dark-grown cells contained 144 ± 11
ribosomes per /i2 of proplastid sectioned, whereas young chloroplasts of 24-h-light
cells (culture B in these experiments) contained 240 ±10 ribosomes per fi2. Fullydeveloped chloroplasts (4 days light) contained 280 ±11 ribosomes per /i 2 of chloroplast sectioned. Thus the actual concentration of ribosomes in the chloroplast approximately doubles during the transition from a proplastid to a fully-developed chloroplast. Since in this transition the proplastid increases fivefold in volume, there is
actually a tenfold increase in the number of chloroplast ribosomes during chloroplast
development in Ochromonas danica.
Chloroplast DNA. In cross-sections through the green chloroplast of Ochromonas
one characteristically sees two DNA areas, one at each end of the chloroplast, located
just inside the two or three outermost lamellar bands which loop around the ends of
the chloroplast (see, for example, the lower section of the chloroplast in Fig. 1). These
DNA areas are approximately 100-250 m/i in cross-section and can be easily recog-
332
S. P. Gibbs
nized even at relatively low magnification because of their low electron density, which
contrasts with the moderate electron density of the rest of the chloroplast matrix. At
higher magnification (Fig. 3) one can distinguish fine strands, approximately 40 A in
diameter, crossing these electron-translucent areas. This appearance is so characteristic
of the nucleoplasm of bacteria and the DNA of mitochondria (Nass & Nass, 1963 a, b)
and chloroplasts (Kislev, Swift & Bogorad, 1965; Gunning, 1965; Bisalputra &
Bisalputra, 1967) that one could reasonably conclude on cytological grounds alone that
these areas contain DNA. However, as additional evidence, greening cells were
labelled with pHJthymidine for 18 h. In autoradiographs prepared from these cells, the
chloroplast grains were preferentially associated with the electron-translucent areas
(Fig. 4). Thus there is little doubt that the peripheral electron-translucent areas contain the DNA of the chloroplast. It should be pointed out that these peripheral DNA
areas are not isolated entities. Since in every cross-section through the plate-like
chloroplast of Ochromonas a DNA area is present in the same location at each extremity,
the DNA in three dimensions can best be interpreted as having the shape of a continuous cord, or ring, which encircles the periphery of the chloroplast. A detailed
description of the three-dimensional configuration of the chloroplast DNA is in
preparation.
Mitochondria. The mitochondria of Ochromonas contain numerous tubular cristae
or villi with electron-translucent lurnina. The mitochondrial matrix is moderately
dense and contains numerous small dense granules, 150-170 A in diameter (Fig. 5).
Ribonuclease-digestion studies of these granules have not been made, but they are
identified as mitochondrial ribosomes on the basis of their size, affinity for lead stains,
and the observations of Andr6 & Marinozzi (1965) and others that similar granules in
the mitochondria of other species are digestible with ribonuclease. In Ochromonas
mitochondria the ribosomes lie both free in the matrix and attached to the villi (arrows,
Fig. 5). Often rows of 3-8 mitochondrial ribosomes are observed attached to the inner
membrane of the mitochondrial envelope. In contrast to the chloroplast ribosomes,
the numbers of mitochondrial ribosomes showed no significant variation in dark- and
light-grown cells.
The only other prominent constituents of the mitochondrial matrix are the striated
cores (Figs. 1 (inset), 5). These are moderately dense areas 120-160 m/i in diameter,
which display parallel longitudinal striations, 40-50 A wide, spaced 50 A apart.
Although DNA is presumably present in these mitochondria, no electron-translucent
areas of fine fibrils have been observed. Possibly the mitochondrial DNA is localized
in the striated cores.
Autoradiographic results
Specificity ofuridine-^fi^H] labelling. Digestion of thin sections of glycol methacrylate-embedded cells with ribonuclease for 30 min reduced the number of grains to
background level. Similar sections exposed 30 min to distilled water alone showed no
reduction in the number of grains per cell. Extraction of the ribonuclease-digested
sections and the distilled water-digested sections with cold 5 % TCA for 1 min did
not significantly alter the grain count in either case. The customary io-min cold TCA
RNA synthesis in chloroplasts and mitochondria
333
rinse was reduced to 1 min in these experiments because it was found that 10 min
extraction removed both the colloidion support film and the glycol methacrylate section. It seems likely, however, that vigorous dipping of an ultrathin section in cold
TCA for 1 min should be sufficient to remove any unbound precursor in the tissue.
Thus it appears that the uridine-s^pH] was specifically incorporated into RNA and
that non-incorporated labelled uridine was removed during the fixing and embedding
procedures.
Percentage distribution of grains after short and long labelling times. Two cultures of
Ochromonas were used in the pHJuridine labelling experiments: one (culture A) had
been exposed to light for 2 h and contained 0-9 x io"10 mg chlorophyll a per cell; the
other (culture B) had been exposed to light for 24 h and contained 12-8 x io~10 mg
chlorophyll a per cell. Aliquots from each of these cultures were labelled with uridine-s^pH] for a relatively short (30 min) and a long (2 h) period in the light before
being prepared for autoradiography. The results of these experiments are summarized
in Tables 1-5 and Figs. 6 and 7.
Table 1 summarizes the percentage distribution of grains over the major cell compartments in each of the 4 experiments. Although background grains make up from 4
to 9% of the grains counted, Tables 4 and 5 show that on an area basis background
labelling is actually very low (only 1 to 2 grains per 100 /i2 of section). Similarly, comparison of Table 1 with Tables 4 and 5 shows that although the grains lying over the
leucosin vacuole constitute a sizeable percentage of the total, when they are expressed
per area of vacuole sectioned, they are not above the level of background labelling.
The leucosin vacuole is a large membrane-limited vacuole which contains the carbohydrate reserve of the organism and it would not be expected to contain any RNA. The
fact that the same level of background labelling was observed outside the cell and over
the leucosin vacuole indicates that there was no chemical fogging of the emulsion by
the fixed cells in these experiments.
Ochromonas damca, in addition to being a photosynthetic and heterotrophic organism,
is also a phagotrophic one, and even when well fed, as in these experiments, often contains one or more food vacuoles with the remains of a partially digested Ochromonas
inside. The cell debris in these food vacuoles is often heavily labelled (Table 1), indicating the rapidity of the capturing and digestive processes, but as these grains are
actually ' exterior' to the cell, they are of no significance in this study and are omitted
from all following calculations.
Thus, the only grains which are of significance in Table 1 are those over the
nucleus and the non-vacuolate cytoplasm. It is interesting that even after only 30 min
of pHJuridine incorporation over three-quarters of the cell grains are extra-nuclear.
This is in contradiction to the earlier light-microscope studies on other organisms of
similar generation time, and the question is whether these extra-nuclear grains are
associated with the mitochondria and chloroplast or with the rest of the cytoplasm.
Table 2 reveals that in the 30-min labelling experiments (A-i and B-i) approximately
two-thirds of the cytoplasmic grains are associated with the mitochondria and chloroplast and only one-third he over the remaining cytoplasm. After 2 h piTJuridine incorporation (experiments A-2 and B-2) there is a marked increase in the proportion of
S. P. Gibbs
334
cytoplasmic grains which are not associated with either the mitochondria or chloroplast, from one-third to one-half of the total. Figures 6 and 7 illustrate this change in
labelling pattern. The cell shown in Fig. 6 has been labelled for 30 min. Both the
nucleolus and chloroplast are well labelled; of the remaining cytoplasmic grains, all
but one lies over a mitochondrion. Figure 7 shows the cytoplasm of a cell which was
labelled with pH]uridine for 2 h. The chloroplast is well labelled, but most of the
other grains lie free in the cytoplasm, not over the mitochondria. This increase in the
proportion of grains associated with the cytoplasmic matrix in cells labelled for 2 h
Table 1. Distribution of total grain counts after uridine-$,6\?lJ] labelling
PreExpt and illumination Labelling
culture no.
time
time
(h)
(h)
Total grain
count
A-1
2
o-5
1081
A-2
B-i
B-2
2
2
1796
93°
1681
24
24
°'5
2
Nucleus
Nonvacuolate
cytoplasm
Food
Leucosin
vacuoles vacuole
12
65
75
75
2
12
73
1
19
13
4
1
Background
6
7
4
5
5
6
9
6
Table 2. Distribution of cytoplasmic grain counts after different times ofuridine-$,6\?H]
incorporation
Expt and
culture no.
Preillumination
time
(h)
A-1
A-2
B-i
B-2
2
2
24
24
Labelling
time
(h)
Total grain
count
Chloroplast
Mitochondria
26
38
29
27
23
698
1297
698
20
42
29
1223
Remaining
cytoplasm
36
51
3i
48
Table 3. Volume of cell occupied by different cell components
Total
time
Expt and
culture no. in light
(h)
A-1
A-2
B-i
B-2
2-5
4
24-5
26
Remainder
of
Leucosin
Food
Nucleolus nucleus vacuole vacuoles
o-5
o-5
o-3
0-3
Chloroplast
Mitochondria
5'9
5'4
1-9
17
6-5
57
2-O
26
48
2-1
5-8
2'I
52
o-8
5-9
1-7
i-8
48
1-2
12
49
i-5
11
Volutin Remaining
granules cytoplasm
i-6
30
31
28
29
suggests that there is a lag in the labelling of the true cytoplasm and that there is not
a corresponding lag in the labelling of the mitochondria and the chloroplast.
Concentration of grain counts over each cell component after short and long labelling
times. In order to determine precisely the comparative rates of labelling of the chloro-
RNA synthesis in chloroplasts and mitochondria
335
plast, mitochondria, and remaining cytoplasm, it was necessary to determine the concentration of grain counts (in grains/unit area) over each cell component at each labelling time. To do this a tracing was made of each micrograph and its component parts
were cut out and weighed as described in Materials and Methods. Since, for each
variable, 400-500 random cell sections were traced, representing over 300 different
Table 4. Grain counts/unit area after different times of uridine-5,6pH] incorporation
by cells pre-illuminated for 2 h (culture A)
30 min labelling
Cell component
Nucleolus
Nucleoplaam
Chloroplast
Mitochondria
Remaining cytoplasm
Leucosin vacuole
Background
(grains/100 /*•)
ISO
32
24
34
6-2
2 h labelling
(grains/ico fi1)
264
48
4i
63
20
i-o
2-2
o-8
1-7
Ratio,*
2 h/30 min
1-8
i-5
1-7
1-9
3'4
—
—
• Before determining ratios, absolute grain counts were corrected for background grains.
Table 5. Grain counts/unit area after different times of uridine-5,6pH] incorporation
by cells pre-illuminated for 24 h (culture B)
30 min labelling
Cell component
Nucleolus
Nucleoplasm
Chloroplast
Mitochondria
Remaining cytoplasm
Leucosin vacuole
Background
(grains/100 /t1)
2 h labelling
(grains/100 /i1)
147
352
37
67
39
55
24
29
7-5
23
i-o
2-O
o-8
2'I
Ratio, •
2 h/30 min
2-4
i-8
i-6
1-9
3-4
—
—
• Before determining ratios, absolute grain counts were corrected for background grains.
cells, this procedure, in addition to giving the total area of each organelle sectioned,
gives the relative volume occupied by each cell organelle in osmium-fixed cells of
Ochromonas. The values obtained are shown in Table 3, even though they are not
necessary for the argument, because relatively few cell organelle volume analyses have
been made from electron micrographs (see Loud, Barany & Pack, 1965).
The grain counts observed per 100 fi% of organelle sectioned are given for the
2-h-light cells (culture A) in Table 4 and for the 24-h-light cells (culture B) in Table 5.
The results for the two cultures are very similar. The chloroplast in culture B is
double the volume of the plastid in culture A, but since the amount of chloroplast
RNA synthesized has increased proportionately, on an area basis, the amount of label
present is the same.
Looking then just at Table 4, it can be seen that after 30 min [3H]uridine incorpora-
336
S. P. Gibbs
tion the nucleolus is very heavily labelled (150 grains/100 /*2) while the nucleoplasm,
the chloroplast, and the mitochondria are all moderately labelled (approximately
30 grains/100 /i2). The remaining cytoplasm, which on a percentage basis contains 29 %
of the total cell grains, is actually only lightly labelled (6 grains/100/i2). It is striking
how much more concentrated the newly labelled RNA is in the mitochondria and
chloroplast than it is in the surrounding cytoplasm. This fact alone suggests that mitochondrial and chloroplast RNA is synthesized in situ and is not transferred to these
organelles via the cytoplasm.
Some indication of the time course of labelling of each organelle is given by the
ratio of the amount of label present in the organelle after 2 h incorporation to that
present after 30 min incorporation. It should be pointed out that for the cell as a
whole the amount of labelling was not proportional to time; rather the total number of
grains present in the cells labelled for 2 h was only 2-1 times the number present at
30 min. This means that either a large proportion of the RNA labelled in 30 min was
short-lived RNA or else the rate of RNA synthesis in the cell as a whole fell off with
time. Calculations showed that in order to explain this ratio in terms of the presence
of a short-lived RNA, over 82 % of the labelled RNA present after 30 min would have
to be the short-lived RNA. (For these calculations, it was assumed that the cell contained a short-lived RNA with a half-life of 20 min or less and a long-lived RNA which
did not turn over at all in 2 h.) This is so unlikely that one must conclude that there
was a fall-off in RNA synthesis with time. This is very probable because log-phase
cells were concentrated 20-fold just prior to the addition of isotope, and it is likely
that some metabolite soon became limiting and the growth rate declined.
If one looks at the ratio observed for each cell component, keeping in mind that a
ratio of 2-1 was observed for the whole cells, it can be seen that there is a marked lag
in the appearance of labelled RNA in the cytoplasm proper, whereas there is no
apparent lag in the appearance of labelled RNA in the mitochondria and chloroplast.
Such a lag in the appearance of labelled RNA in the cytoplasm is in line with present
knowledge that this RNA is made in the nucleus. The absence of such a lag in the
labelling of the mitochondria and chloroplast adds further evidence that the RNA of
these organelles is not transferred from the nucleus, but is made in situ.
DISCUSSION
The above autoradiographic observations meet the prediction that organelles
capable of RNA synthesis should show a high level of uridine incorporation and an
independence of cytoplasmic labelling. To this extent, the results provide support
from yet another field for a degree of autonomy in chloroplasts and mitochondria,
which has long been suspected on genetical, physiological and, more recently, biochemical grounds.
It should be pointed out that the evidence presented above for in situ RNA synthesis is not in itself conclusive, since other interpretations are at least conceivable, for
instance that nuclear-synthesized RNA is preferentially directed to the organelles
when they are in active development. Fortunately, in the course of this work, a more
RNA synthesis in chloroplasts and mitochondria
337
direct confirmation of the independent synthesis of chloroplast RNA was provided by
the observation that chloroplast RNA is synthesized at the site of the chloroplast
DNA (Gibbs, 1967). Since chloroplast DNA in greening cells of Ochromonas has a
restricted and easily identified location at the periphery of the chloroplast, it was
possible to show that after 30 min exposure to uridine-s^pH] the chloroplast label
was concentrated near this peripheral DNA ring, whereas after 2 h incorporation it
had become uniformly distributed throughout the chloroplast (Gibbs, 1967).
Of major interest is the question of which kinds of RNA are synthesized within the
chloroplast. These experiments cannot, of course, give a direct answer. However,
since chloroplast ribosomes increase in number tenfold during chloroplast development in Ochromonas, and since in other cells with labelling times as long as 30 min the
bulk of the RNA synthesized is ribosomal RNA, one can conclude with reasonable
certainty that chloroplast ribosomal RNA is synthesized within the chloroplast of
Ochromonas. This conclusion receives support from Berger's (1967) observation that
ribosomal RNA is synthesized in chloroplasts isolated from previously enucleated
plants of Acetabularia and from Scott & Smillie's (1967) recent evidence that chloroplast ribosomal RNA from Euglena hybridizes with the chloroplast DNA. In light of
Berger's (1967) report that 4 s RNA is synthesized in isolated chloroplasts of Acetabularia, it seems likely that transfer RNA is also synthesized within the chloroplast
of Ochromonas. The vital question of whether chloroplast messenger RNA's are synthesized in situ must await more sophisticated studies.
The three sizes of ribosomes observed in this study—210-230 A ribosomes in the
cytoplasm, 170-200 A ribosomes in the chloroplast, and 150-170 A ribosomes in the
mitochondria—are in accord with the different sedimentation coefficients reported for
the three types of ribosomes in other species. Lyttleton (1962), Boardman et ah (1966),
and others have observed that in higher plants, in general, chloroplast ribosomes have
a lower sedimentation coefficient (70 s) than cytoplasmic ribosomes (80 s), and recently
O'Brien & Kalf (1967a, b) have reported that the ribosomes from rat liver mitochondria have a sedimentation coefficient of 55 S.
I am indebted to Professor C. H. Waddington for his support and encouragement. I also
wish to thank Dr J. Jacob for his skilful assistance and advice and Dr J. L. Sirlin and Dr R. J.
Poole for many valuable discussions. This investigation was supported by the Medical Research
Council of Great Britain and the National Research Council of Canada (Grant no. A-2921).
REFERENCES
AARONSON, S. & BAKER, H. (1959). A comparative biochemical study of two species of Ochromonas. J. Protozool. 6, 282-284.
ANDRE, J. & MARINOZZI, V. (1965). Presence, dans les mitochondxies, de particules ressemblant
aux ribosomes. J. Microscopie 4, 615-626.
BARNETT, W. E. & BROWN, D. H. (1967). Mitochondrial transfer ribonucleic acids. Proc.
natn. Acad. Set. U.S.A. 57, 452-458.
BERGER, S. (1967). RNA-synthesis in Acetabularia. II. RNA-synthesis in isolated chloroplasts.
Protoplasma 64, 13-25.
BISALPUTRA, T. & BISALPUTRA, A. (1967). The occurrence of DNA fibrils in chloroplasts of
Lmirencia spectabilis. J. Ultrastruct. Res. 17, 14-22.
338
S. P. Gibbs
N. K., FRANCKI, R. I. B. & WILDMAN, S. G. (1966). Protein synthesis by cell-free
extracts of tobacco leaves. III. Comparison of the physical properties and protein synthesizing
activities of 70 S chloroplast and 80 S cytoplasmic ribosomes. J. molec. Biol. 17, 470-489.
BRAWERMAN, G. & EISENSTADT, J. M. (1964). Template and riboaomal ribonucleic acids associated with the chloroplasts and the cytoplasm of Euglena gracilis. J. molec. Biol.
BOARDMAN,
10,403-411.
L. G. & TUBERGEN, R. P. VAN (1962). High-resolution autoradiography. I. Methods.
J. Cell Biol. 15, 173-188.
CHAYES, F. (1965). Determination of relative volume by sectional analysis. Lab. Invest. 14,
987-1008.
CHEN, J. L. & WILDMAN, S. G. (1967). Functional chloroplast polyribosomes from tobacco
leaves. Science, N.Y. 155, 1271-1273.
CHUN, E. H. L., VAUGHAN, M. H. Jr. & RICH, A. (1963). The isolation and characterization of
DNA associated with chloroplast preparations. .7. molec. Biol. 7, 130-141.
CLARK, M. F. (1964). Polyribosomes from chloroplasts. Biochim. biophys. Acta 91, 671-674.
CLARK, M. F., MATHEWS, R. E. F. & RALPH, R. K. (1964). Ribosomes and polyribosomes in
Brassica pekinensis. Biochim. biophys. Acta 91, 289—304.
GIBBS, S. P. (1962). Chloroplast development in Ochromonas danica.J. Cell Biol. 15, 343-361.
GIBBS, S. P. (1967). Synthesis of chloroplast RNA at the site of chloroplast DNA. Biochem.
biophys. Res. Cowman. 28, 653-657.
GRANICK, S. & GIBOR, A. (1967). The DNA of chloroplasts, mitochondria, and centrioles. In
Progress in Nucleic Acid Research and Molecular Biology, vol. 6 (ed. J. N. Davidson & W. E.
Cohn), pp. 143-186. New York and London: Academic Press.
GUNNING, B. E. S. (1965). The fine structure of chloroplast stroma following aldehyde osmiumtetroxide fixation. J. Cell Biol. 24, 79-93.
IWAMURA, T. (1966). Nucleic acids in chloroplast and metabolic DNA. In Progress in Nucleic
Acid Research and Molecular Biology, vol. 5 (ed. J. N. Davidson & W. E. Cohn), pp. 133155. New York and London: Academic Press.
JACOBSON, A. B., SWIFT, H. & BOGORAD, L. (1963). Cytochemical studies concerning the
occurrence and distribution of RNA in plastids of Zea mays. J. Cell Biol. 17, 557-570.
JANOWSKI, M. (1965). Synthese chloroplastique d'acides nucleiques chez Acetabularia mediterranea. Biochim. biophys. Acta 103, 399-408.
KALF, G. F. (1964). Deoxyribonucleic acid in mitochondria and its role in protein synthesis.
Biochemistry 3, 1702-1712.
KIRK, J. T. O. (1964). DNA-dependent RNA synthesis in chloroplast preparations. Biochem.
biophys. Res. Contmun. 14, 393-397.
KISLEV, N., SWIFT, H. & BOGORAD, L. (1965). Nucleic acids of chloroplasts and mitochondria
in Swiss chard. J. Cell Biol. 25, 327-344.
LEDUC, E. H., BERNHARD, W. & TOURNIER, P. (1966). Cyclic appearance of atypical mitochondria containing DNA fibers in cultures of an adenovirus 12-induced hamster tumor. Expl
Cell Res. 43, 597-616.
LEDUC, E., MARINOZZI, V. & BERNHARD, W. (1963). The use of water-soluble glycol methacrylate in ultrastructural cytochemistry. Jl R. microsc. Soc. 81, 119-130.
LOUD, A. V., BARANY, W. C. & PACK, B. A. (1965). Quantitative evaluation of cytoplasmic
structures in electron micrographs. Lab. Invest. 14, 996—1008.
LUCK, D. J. L. & REICH, E. (1964). DNA in mitochondria of Neurospora crassa. Proc. natn.
Acad. Sci. U.S.A. 52, 931-938.
LYTTLETON, J. W. 1962. Isolation of ribosomes from spinach chloroplasts. Expl Cell Res. 26,
CARO,
312-317.
M. M. K. & NASS, S. (1963 a). Intramitochondrial fibers with DNA characteristics. I.
Fixation and electron staining reactions. J. Cell Biol. 19, 593-611.
NASS, S. & NASS, M. M. K. (19636). Intramitochondrial fibers with DNA characteristics. II.
Enzymatic and other hydrolytic treatments. J. Cell Biol. 19, 613-629.
NEUBERT, D. & HELGE, H. (1965). Studies on nucleotide incorporation into mitochondrial RNA.
Biochem. biophys. Res. Contmun. 18, 600-605.
NEWCOMB, E. H. (1967). Fine structure of protein-storing plastids in bean root tips. J. Cell
Biol. 33, 143-163.
NASS,
RNA synthesis in chloroplasts and mitochondria
339
T. W. & KALF, G. F. (1967a). Ribosomes from rat liver mitochondria. I. Isolation
procedure and contamination studies.^, biol. Chem. 242, 2172-2179.
O'BRIEN, T. W. & KALF, G. F. (19676). Ribosomes from rat liver mitochondria. II. Partial
characterization. J. biol. Chem. 242, 2180-2185.
PRESCOTT, D. M. (1964). Cellular sites of RNA synthesis. In Progress in Nucleic Acid Research
and Molecular Biology, vol. 3 (ed. J. N. Davidson & W. E. Cohn), pp. 33-57. New York and
London: Academic Press.
PRINGSHEIM, E. G. (1955). Kleine Mitteilungen iiber Flagellaten und Algen. III. Ober Ochromonas danica n.sp. und andere Arten der Gattung. Arch. Mikrobiol. 23, 181-192.
REYNOLDS, E. S. (1963). The use of lead citrate at high pH as an electron-opaque stain in
electron microscopy.^. Cell Biol. 17, 208-213.
Ris, H. & PLAUT, W. (1962). Ultrastructure of DNA-containing areas in the chloroplast of
Chlamydomonas. J. Cell Biol. 13, 383-391.
SCHUSTER, F. L. (1965). A deoxyribose nucleic acid component in mitochondria of Didyntium
nigripes, a slime mold. Expl Cell Res. 39, 329-345.
SCHWEIGER, H. G. & BERGER, S. (1964). DNA-dependent RNA synthesis in chloroplasts of
Acetabularia. Biochim. biophys. Acta 87, 533-535.
SCHWEIGER, H. G., DILLARD, W. L., GIBOR, A. & BERGER, S. (1967). RNA-synthesis in
Acetabularia. I. RNA-synthesis in enucleated cells. Protoplasma 64, 1-12.
SCOTT, N. S. & SMILLIE, R. M. (1967). Evidence for the direction of chloroplast ribosomal
RNA synthesis by chloroplast DNA. Biochem. biophys. Res. Commun. 28, 598-603.
SHAH, V. C. & LYMAN, H. (1966). DNA-dependent RNA synthesis in chloroplasts of Euglena
gracilis. J. Cell Biol. 29, 174-176.
SHEPHARD, D. C. (1965). An autoradiographic comparison of the effects of enucleation and
actinomycin D on the incorporation of nucleic acid and protein precursors by Acetabularia
chloroplasts. Biochim. biophys. Acta 108, 635-643.
SISSAKIAN, N. M., FILIPPOVICH, I. I., SVETAILO, E. N. & ALIYEV, K. A. (1965). On the proteinsynthesizing system of chloroplasts. Biochim. biophys. Acta 95, 474-485.
SPENCER, D. (1965). Protein synthesis by isolated spinach chloroplasts. Archs Biochem. Biophys.
i n , 381-390.
SPENCER, D. & WHITFELD, P. R. (1966). The nature of the ribonucleic acid of isolated
chloroplasts. Archs Biochem. Biophys. 117, 337-346.
STUTZ, E. & NOLL, H. (1967). Characterization of cytoplasmic and chloroplast polysomes in
plants; Evidence for three classes of ribosomal RNA in nature. Proc. natn. Acad. Sci. U.S.A.
57, 774-781.
SWIFT, H., ADAMS, B. J. & LARSEN, K. (1964). Electron microscope cytochemistry of nucleic
acids in Drosophila salivary glands and Tetrakymena. jfl R. microsc. Soc. 83, 161-167.
WINTERSBERGER, E. (1965). Proteinsynthese in isolierten Hefe-Mitochondrien. Biochem. Z. 341,
409-419.
WINTERSBERGER, E. (1966). Occurrence of a DNA-polymerase in isolated yeast mitochondria.
O'BRIEN,
Biochem. biophys. Res. Commun. 25, 1-7.
E. & TUPPY, H. (1965). DNA-abhangige RNA-Synthese in isolierten HefeMitochondrien. Biochem. Z. 341, 399-408.
WINTERSBERGER,
{Received 6 October 1967)
340
S. P. Gibbs
Fig. i. Longitudinal section of a 20-h-light cell of Ochromonas danica. Since the platelike chloroplast of Ochromonas is bent in the shape of a U around the nucleus, the
chloroplast has been sectioned twice. An electron-translucent DNA area (dr) can be
seen at each extremity of the lower section and at the posterior end of the upper section.
In three dimensions, the chloroplast DNA has the shape of a ring which encircles the
periphery of the chloroplast. Abundant chloroplast ribosomes (cr) and larger osmiophilic granules (og) are present in the chloroplast matrix. Nuclear pores (np) are present
in the nuclear envelope where it faces the cytoplasm, but are absent from the regions of
the nuclear envelope (between arrows) which form one wall of the sac of endoplasmic
reticulum (cer) enclosing the chloroplast. (Iv, leucosin vacuole.) x 35000. Inset:
mitochondrion with a prominent striated core (sc). x 48000.
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S. P. GIBBS
(Facing p. 340)
Fig. 2. Proplastid of a dark-grown cell. Scattered chloroplast ribosomes (cr) and a
single lamellar disc (d) are visible, (cer, chloroplast ER.) x 48000.
Fig. 3. Chloroplast of a 20-h-light cell. Fine fibrils can be seen in the peripheral DNA
area (arrow), (cer, chloroplast ER; cr, chloroplast ribosomes; n, nucleus.) x 48000.
Fig. 4. Autoradiograph of a greening cell of Ocliromonas danica labelled with [3H]thymidine for 18 h. In each lobe of the chloroplast the peripheral DNA ring has been transected twice. Three out of the 4 DNA areas are labelled (arrows). The unlabelled DNA
area (dr) is partially obscured by dirt in this preparation, x 16000.
Journal of Cell Science, Vol. 3, No. 3
• V •
S. P. GIBBS
Fig. 5. Grazing section of a 4-day-light cell illustrating the mitochondria and microtubules (mt) particularly well. Mitochondrial ribosomes are present both in the matrix
(mr) and attached to the tubular cristae (arrows). Striated cores (sc) are sectioned in
most of the mitochondria, but it is difficult to resolve their fine structure in this
micrograph, (c, chloroplast.) x 48000.
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S. P. GIBBS
Fig. 6. Autoradiograph of a 2-h-light cell labelled with uridine-5,6['H] for 30 min. In
this cell, all but one of the cytoplasmic grains are associated with mitochondria or the
chloroplast. x 16000.
Fig. 7. Autoradiograph of a 24-h-light cell labelled with uridine-5,6[3H] for 2 h. The
majority of the cytoplasmic grains in this cell lie over the background cytoplasm,
x 16000.
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S. P. GIBBS