J. Cell Set. 36, 281-299 (1977)
Printed in Great Britain
281
VISUALIZATION OF INTERPHASE
CHROMOSOMES
STEPHEN M. STACK,• DAVID B. BROWN,* AND
WILLIAM C. DEWEYf
Department of Botany and Plant Biology* and Department of Radiology and Radiation
Biologyf, Colorado State University, Fort Collins, Colorado 80523, U.S.A.
SUMMARY
Using a modified Giemsa-banding technique we have observed what appear to be chromosomes during interphase in nuclei from Allium cepa root tips and Chinese hamster cells (CHO
line). During telophase through Gx chromosomes progressively uncoil and decondense.
During S chromosomes are comparatively decondensed, but some segments have structure
similar to chromosomes in. GL and G,. During Gt the chromosomes progressively recondense
and coil in apparent preparation for prophase. Although specific structural modifications of
chromosomes occur in Glt S, and Gx nuclei, chromosomes appear never to decondense to the
point that they lose their 3-dimensional integrity, but remain in distinct domains throughout
interphase.
INTRODUCTION
Chromosomes typically are visible during cell division in eukaryotes, but during
telophase chromosomes usually become indistinct in a process referred to as decondensation (swelling) and/or uncoiling (despiralization or unravelling) (Brown &
Bertke, 1974). When chromosomes have disappeared in the interphase nucleus,
usually there is no clear indication of chromosome boundaries or even at what level
the structural integrity of chromosomes is maintained to allow them to reappear in
subsequent divisions. Current models for the structure of interphase nuclei assume
an unravelling of chromosomes into chromonemata that mingle in the nucleoplasm in
the form of fibres 10-30 nm in diameter (Comings, 1968; DuPraw, 1970; Brasch &
Setterfield, 1974). The structural integrity of interphase chromosomes is thought to
depend on numerous attachments of chromosomal fibres to the nuclear envelope.
These models are based largely on electron microscopy of sectioned and wholemounted interphase nuclei in which usually only a tangle of chromonemata can be
observed, with no sign of individual chromosomes (Wischnitzer, 1973).
In apparent contradiction to this model for interphase chromosomes, we have
observed what seem to be distinct interphase chromosomes throughout the cell cycle of
Allium cepa and Chinese hamster (CHO line) nuclei that have been treated with a modification of the chromosome banding technique of Drets & Shaw (1971). Our observations
are generally in agreement with many older light-microscopic descriptions of interphase
nuclei and chromosomes, and we interpret these observations as being compatible with
and complementary to descriptions of interphase nuclei based on electron microscopy.
282
S. M. Stack, D. B. Brown and W. C. Dewey
MATERIALS AND METHODS
Allium cepa bulbs were suspended in aerated tap water at room temperature until roots sprouted.
The bulbs were then transferred to damp vermiculite until the roots grew to a length of 2-5 cm.
At this time the roots were either cut from the bulbs or allowed to grow one hour longer in
distilled water containing 10 /iCi/ml tritiated thymidine (New England Nuclear, 20 Ci/mmol)
before being cut from the bulbs. Harvested roots were immediately fixed for 1 h in aceticethanol (1:3). Using 1 or 2 root tips in a drop of 45 % acetic acid on a slide, meristematic cells
were picked out with dissecting needles, and the multicellular fragments were removed. These
essentially single cell suspensions were squashed and the coverslips removed by the dry-ice
method. After the slides were air-dried briefly, they were stored in a CaClj desiccator foi
4-5 h. Dried slides were then treated according to one of the schedules (I-V) in Table 1.
Although many variations of schedules I I-V were tried and some were reasonably effective in
showing structure in interphase nuclei, the exact schedules in treatments I I-V were generally
superior.
Chinese hamster cells (CHO line) were maintained in logarithmic growth in McCoy's
5A medium containing 15 % foetal calf serum. Cells were synchronized in mitosis by shaking
off mitotic cells, after which separate flasks were harvested by txypsinization at o, 1, 3, 6, 10,
and 12 h of incubation (Dewey, Noel & Dettor, 1972). Immediately before trypsinization, the
cells were pulse labelled with 5 /tCi/ml tritiated thymidine for 15 min. Trypsinized cells were
fixed for 5 min in 1:3 acetic ethanol and then squashed immediately in 50% acetic acid.
Coverslips were removed by the dry-ice method, and the squashes were air dried for 1-9 days
in open slide boxes. Dried slides were then treated according to schedules I, II, or IV in Table 1.
At o h the cells were 95 % mitotic. After 1 h, none of 200 nuclei scored were labelled, indicating
nuclei were exclusively in Gx. After 3 h only 7 % of the nuclei were labelled, so most cells were
in late G1 while a few cells had passed into S. After 6 h 72 % of the nuclei were labelled, so by
this time most cells were in early S phase with the remainder in late Gx. After 10 h 96 % of the
nuclei were labelled, indicating most cells were in middle to late 5. After 12 h 41 % of the
nuclei were labelled, and mitotic cells were frequently observed. Apparently by this time
synchrony had been largely lost, since many cells had passed into and through G, while the
remainder were in late S (Dewey et al. 1972).
For autoradiography, dried slides were dipped in Kodak NTB2 liquid emulsion at 41 °C,
air dried overnight in a light-proof box with air circulating through it, and stored 4-6 days in
a sealed light-proof box at 4 °C. The slides were developed 2 min in Kodak D19 developer at
20 CC, rinsed in distilled water, and fixed 3 min in Kodak (acid) fixer. After washing 5-10 min
in distilled water, the slides were air dried before further processing.
Giemsa staining was performed by flooding the slides with a 1:9 mixture of ice-cold 620
Harleco stock Giemsa stain and 0-12 M, pH 68 potassium phosphate buffer. After staining
20 min, the slides were rinsed in distilled water and air dried.
Although nuclei treated for autoradiography (see above) can show structured interphase
chromatin reasonably well, the clearest demonstrations are shown by nuclei that have not been
treated for autoradiography. To combine a good demonstration of interphase structure with
tritiated thymidine labelling, schedule IV in Table 1 involves first treating nuclei to produce
interphase structure followed by photography, destaining, treating for autoradiography, and
restaining. Although this procedure sometimes worked reasonably well, there was always a
question, particularly in A. cepa, whether the Giemsa stain had been sufficiently washed out
before the treatment for autoradiography. If any Giemsa stain remains, it somehow exposes
the emulsion regardless of the presence or absence of tritium (see Acknowledgements). For
this reason, this method was used only to a limited extent for A. cepa, but it was used extensively
for CHO cells where it seemed easier to wash out the Giemsa stain adequately.
All light microscopy and photomicrography were performed on a Leitz Orthoplan microscope
fitted with a 4* x 5* format automatic camera using Kodak Ektapan film, or a 35-mm camera
using Kodak Panatomic X film.
Interphase stages (G1( S and G,) in A. cepa nuclei were determined by pulse labelling with
tritiated thymidine and from differences in nuclear diameters. Nuclear diameters were assumed
to be positively correlated with nuclear volumes that generally have been shown to be pro-
Visualization of interphase chromosomes
283
portional to DNA content in both plants (Sunderland & McLeish, 1961) and animals (Sawicki,
Rowinski & Swenson, 1974). On this basis the smallest nuclei were considered to be in Glt
intermediate size nuclei were considered to be in S, and the largest nuclei were considered to
be in Gt. Although the correlation of DNA content with nuclear volume seems generally reliable for rapidly dividing cells, slowly dividing and non-mitotic cells are often exceptional
Table 1. Flaw chart outlining various treatment schedules (I—V) of dried squashes on
slides. All schedules start at the top of the chart with dried slides and end at the bottom
of a column indicated by a Roman numeral for the particular schedule
dried slides
i
stain in 2 % acetoorcein for io min
submerge in a solution that is
002 N NaOH and 0-114 M NaCl at
22 °C for io s
4
wash off coverslip
and staining solution
with ioo % ethanol and
air dry
4
3 rapid washes in 70 % ethanol
4
3 rapid washes in 95 % ethanol
i
1
treat for autoradiography
(see Materials
and methods)
1
{
mount in
Euparal
I
1
mount in
Euparal
1
I
4
photograph (see
Materials and
methods)
1
1
wash off coverslip and Eupaial
with ioo% ethanol
and air dry
4
treat air-dried
slides according
to schedule three
II
submerge in 2 x SSC (Drets & Shaw,
1971) at 60 °C for 1 min
1
1.
Giemsa stain
(see Materials
and methods)
I
flood with icecold O-I2 M, pH
68 potassium phosphate
huflrer flnrl Allow to
1
1
4
photo- photograph graph
III
/
/
stand for 10 min
1
1
rinse in distilled
water and air dry
remove coverslip,
Euparal, and
Giemsa stain with
methanol and
ethanol washes
and air dry
4
treat for autoradiography
1
4
Giemsa stain
4
treat for autoradiography
Giemsa stain
mount in
Euparal
4
V
1
4
mount in Euparal
IV
(Macleod & MacLachlan, 1974; Leuchtenberger & Schrader, 1951). To avoid the problem of
contaminating rapidly dividing meristematic cells with the slowly or non-di aiding cells of the
root cap, apical meristem, and zone of elongation, these parts of the root tip were largely cut
away and only a section approximately 2 mm long containing the primary meristems was used to
prepare squashes. The single cells dissected from this section were assumed to be cycling
284
S. M. Stack, D. B. Brown and W. C. Dewey
rapidly and to show the correlation between nuclear volume and DNA content (Sunderland &
McLeish, 1961). CHO cells were harvested only from non-confluent cultures in their log phase
of growth to assure rapidly cycling cells.
Because diameter is also correlated positively with the amount of pressure applied during
squashing, it is necessary to look at cells that have been comparably squashed for accuracy in
estimating interphase stages by nuclear diameters. Although this problem initially seems almost
insurmountable because of the variation in pressure applied from one thumb squash to another,
the solution appears to be intrinsic to the method of visualizing structured interphase chromatin.
In a thumb squash on an 18- or 22-mm square coverslip, there is an area of high pressure in
the centre that integrates with comparatively low pressure at the periphery where the thumb
is not in contact. The central nuclei are usually over-squashed, in the sense that nuclear diameters are very large and no internal structure is visible. However, more peripherally there is
a ring of generally intermediate-size nuclei that surround the central group, and it is here that
nuclei with structured chromatin can be observed. More peripherally, the nuclei are generally
small, dark-staining, and without apparent internal structure. Presumably in this most peripheral
zone the pressure was insufficient to flatten nuclei to the extent necessary to reveal internal
structure. All measurements and photography were concentrated in the intermediate zone
where the pressure was most appropriate. Using this population of cells that varied to some
extent from slide to slide in its precise distance from the centre of the squash, a good correlation
between slides was observed, as diameters of hundreds of nuclei were related to classifications
in Gu S, and Gz.
For electron microscopy, root tips of A. cepa were fixed in phosphate-buffered glutaraldehyde
and osmium tetroxide and embedded in Epon-Araldite (see Packard & Stack, 1976, for details).
Embedded root tips were both thick (i-/im) and thin (80-nm) sectioned on an LKB 4801A
ultramicrotome. Thin sections were stained with uranyl acetate and lead citrate and photographed in an AEI EM6b electron microscope. Thick sections were mounted on glass slides,
stained with 1 % methylene blue in 012 M dibasic potassium phosphate, and photographed light
microscopically. CHO cells were fixed in Karnovsky's fixative and embedded in Epon. Thin
sections were post-stained with phosphotungstic acid and photographed in a Philips 200
electron microscope.
RESULTS
When A. cepa root tip cells or Chinese hamster (CHO line) cells in interphase were
treated by one of schedules II-V in Table 1 (called sodium hydroxide treatments
subsequently), chromatin could be seen condensed into distinct, separate masses that
often resembled chromosomes.
To determine whether chromatin of A. cepa differs in appearance during the various
stages of interphase, nuclei were identified as being in Glt S, or G2 by tritiated thymidine pulse labelling and by comparisons of the diameters of the squashed nuclei (see
Materials and methods). Using these techniques we have arranged the unsynchronized,
sodium-hydroxide-treated nuclei from A. cepa root tips in a sequence that represents
a light-microscopic interpretation of the changes that chromatin undergoes through
the mitotic cycle. A familiar point at which to begin such a sequence is metaphase.
At this time the condensed chromosomes are scattered over the metaphase plate,
indicating A. cepa does not have a hollow spindle (Fig. 1). Often it can be seen that
the chromosomes have heterochromatic telomeres. In anaphase the chromosomes take
on a pole-to-pole orientation with the heterochromatic telomeres trailing to the inside
of each group of separating sister chromosomes (Fig. 2). During telophase the nuclear
envelope forms around the chromosomes as they maintain their relic anaphase
Visualization of interphase chromosomes
285
orientation, and the chromosomes begin to swell (Fig. 3).
Gx follows telophase. Because the cells were not synchronized, nuclei could not be
classified as being in early, middle or late Gx by this means. However, assuming a
progressive decondensation of chromatin during Gx as is generally reported (Taylor,
1963; Lafontaine & Lord, 1974; de la Torre, Sacristan-Garate & Navarrete, 1975), one
can arrange Gx nuclei in a series that appears to show progressive decondensation of
chromosomes that retain their relic anaphase-telophase orientation (Heitz, 1932;
Stack & Clarke, 1974; Fussell, 1975) (Figs. 4-9). Also, the chromosome-like structures
often appear to have heterochromatic telomeres (Figs. 4, 6, 9) like chromosomes from
prophase through anaphase (Figs. 1, 2, 20, 21).
Nuclei in early S phase resemble those interpreted to be late Gx nuclei (Figs. 10—11).
As the cell proceeds through S phase the chromatin becomes increasingly diffuse, but
there are usually some clear divisions between chromatin masses (Figs. 12, 13). If
the cells are squashed intensely, long strands of finely dispersed chromatin are evident
(Fig. 14). These strands probably correspond to chromonemata of linearly intact
chromosomes whose 3-dimensional structure has been destroyed (Rohme, 1975). Late
in S phase, structure again becomes more obvious as nuclei begin to resemble G2
nuclei (Fig. 15).
G2 nuclei have highly structured chromatin that often resembles long chromosomes.
Although cells cannot be classified as early, middle, or late G2 on the basis of synchrony,
G2 nuclei can be arranged in an order that appears to show the progressive condensation of chromosomes for prophase (Figs. 16-19). This order is supported by the
observation that large, lightly labelled nuclei (Fig. 15) that are interpreted to be in
early G2 or late in S phase have less distinct chromosomes than large, unlabelled
nuclei (Figs. 16-18) that are interpreted to be in middle to late G2 (see Materials and
methods). Except for occasional heterochromatic telomeres (Fig. 17), G2 chromosomes
do not appear to be double, but since this technique does not show sister chromatids
at prophase or metaphase (Figs. 1,21), this is not surprising. The condensing chromosomes in late G2 or early prophase vary in diameter apparently because condensation
or coiling does not proceed at the same rate even within individual chromosomes
(Figs. 19, 20). Finally in middle to late prophase, all of the chromosomes appear to
have the same diameter and retain their relic anaphase-telophase orientation
(Fig. 21).
Aceto-orcein-stained nuclei were prepared (Table 1, schedule I) to'compare their
more familiar appearance with sodium hydroxide-treated nuclei. Small unlabelled
telophase nuclei (Fig. 22) typically show chromosome structure that continues
through early Gx (Fig. 23), but structure seems largely to be lost by late Gx (Fig. 23).
During S the nuclei increase in size but have little recognizable structure (Figs. 24,
25). During G2 some thin strands and clumps of chromatin are visible, but these are
difficult to interpret as chromosomes (Figs. 26, 27). By early prophase distinct
thickened strands are visible in relic anaphase-telophase orientation (Fig. 28). As
condensation continues toward middle prophase the strands retain their orientation
and become thinner, with more even diameters (Fig. 29). From these observations it
appears that chromosomes look similar after aceto-orcein staining and sodium
19
CEL
26
286
S. M. Stack, D. B. Brown and W. C. Dewey
16
Visualisation of interphase chromosomes
287
hydroxide treatment from early prophase through early G1( but during most of interphase aceto-orcein staining provides little evidence for the persistence of chromosome
structure.
Nuclei of synchronized CHO cells were identified as being Gx, S or G2 according to
(1) the incubation time, (2) whether they were labelled with tritiated thymidine, and
(3) by comparing nuclear diameters. In the following description of changes in the
structure of sodium hydroxide-treated (schedule IV in Table 1) nuclei during the
cell cycle, all of these means of determining stage were in general agreement.
In the o-h sample, most nuclei were in metaphase (Fig. 30). The chromosomes are
arranged at the periphery of the metaphase plate, which indicates the presence of a
' hollow spindle' that is characteristic of many animals (DuPraw, 1970). This arrangement is carried through anaphase-telophase, with the chromosome arms radiating
from an apparently empty polar area (Fig. 31). In the o-h sample, there were also a
few telophase nuclei that showed what appeared to be progressive stages in the uncoiling and swelling of chromosomes (Figs. 32, 33).
Figs. 1-29. Chromosomes and nuclei horn A. cepa root tips, x 1200. Figs. 1-21 represent sodium hydroxide-treated preparations. The clearest of these were not treated
for autoradiography (see Materials and methods). The Roman numerals in parentheses
that precede the descriptions of each figure indicate which of the schedules in Table 1
was used to make the preparation. Schedule V involves autoradiography while
schedule III does not. Fig. 1: (III) Polar view of a metaphase chromosome spread.
Note the heterochromatic telomeres. Chromosomes are scattered over the metaphase
plate indicating the absence of a hollow spindle. Fig. 2: (III) Lateral view of late
anaphase or early telophase. Again note the heterochromatic telomeres and the
orientation of chromosomes. Fig. 3: (V) Later al view of telophase showing continued
orientation of chromosomes. Fig. 4: (III) Polar view of an early G^ or late telophase
nucleus. Observe the radiating chromosomes with heterochromatic telomeres (arrows).
Fig. 5 : (V) Polar view of a more advanced Gx nucleus with radiating chromosomes that
are swollen compared to Fig. 4. Fig. 6: (III) Polar view of a Gl nucleus that is quite
similar to the nucleus in Fig. 5 except that the radiating chromosomes are clearer and
show heterochromatic telomeres (arrows). Fig. 7: (V) Lateral view of Gx nucleus with
swollen chromosomes visible on one side (large arrow). Fig. 8: (III) Lateral view of
Gi nucleus that is very similar to the nucleus in Fig. 7 except the oriented chromosomes are much clearer. Fig. 9: (III) Lateral view of a late Gj 01 early S-phase nucleus
with oriented chromosomes and heterochromatic telomeres. Chromosomes generally
are becoming less distinct. Fig. 10: (V) Lightly labelled early S-phase nucleus in which
chromosomes are less clear, but divisions still separate chromatin domains that probably
represent chromosomes. Fig. 11: (III) Early S-phase nucleus comparable to the
nucleus in Fig. 10. Heterochromatic telomeres continue to be visible on some
chromosomes. Fig. 12: (V) Labelled mid- to late S-phase nucleus still showing some
separate chromatin domains, but in general such nuclei have more diffuse chromatin
than Gx nuclei. Fig. 13: (III) Mid- to late 5-phase nucleus comparable to the nucleua
in Fig. 12, but chromatin domains are more distinct. Fig. 14: (IV) Labelled 5-phase
nucleus that was over-squashed to reveal a complex network of threads that probably
represent chromonemata that were forced out of their domains. Autoradiographic
emulsion was applied after this photograph was taken. Fig. 15: (V) Lightly labelled
late S or early Gt nucleus with more distinct chromatin domains. Fig. 16: (V) Early
G, nucleus with distinct domains of chromatin.
rg-2
288
S. M. Stack, D. B. Brown and W. C. Dewey
29
Visualization of interphase chromosomes
289
Cells in Gx were studied in the 1- and 3-h samples. In early G± nuclei, swelling
(uncoiling) of the chromosomes can be observed to occur asynchronously or unevenly
along the length of individual chromosomes (Fig. 34). In late Gx nuclei more finely
divided chromatin masses, some of which still seem recognizable as chromosomes, can
be observed (Fig. 35).
In the 6-h sample, early 5 phase nuclei had both chromosomes in the process of
uncoiling and finely divided chromatin that may represent uncoiled chromosomes
(Figs. 36-39). Other nuclei that were interpreted as being later in S phase showed a
combination of finely divided and clumped chromatin (Figs. 40, 41).
In the 10-h sample, lightly labelled nuclei were considered to be very late in S phase
or early in G2. These nuclei showed considerable internal structure that often resembled what could be interpreted as condensing chromosomes (Figs. 42, 43).
Finally, in the 12-h sample, chromosome condensation was underway. Chromosomes appeared to condense (perhaps coil) asynchronously along their length (Figs. 44,
45), and this condensation continued until the chromosomes became very thin by
early prophase (Fig. 46). As prophase continued, chromosomes became thicker and
shorter (Fig. 47) until they reached their metaphase length (Fig. 30).
Aceto-orcein-stained CHO nuclei were prepared to compare their structure to
sodium-hydroxide-treated nuclei. After aceto-orcein staining, G± and G2 nuclei
contained generally diffuse chromatin with some clumping evident (Figs. 48, 50).
S phase nuclei consistently contained more diffuse chromatin (Fig. 49).
Since aceto-orcein staining did not seem to represent a treatment that would
necessarily interfere with subsequent sodium hydroxide treatment to show structured
interphase chromosomes, we thought it might be possible to make a direct comparison
Fig. 17: (III) Two early G% nuclei similar to the nucleus in Fig. 16, but the chromatin domains have more the appearance of separate chromosomes with heterochromatic telomeres, one of which appears double (arrow). Fig. 18: (V) Lateral view
of a Ga nucleus in which there is a suggestion of oriented chromosomes. Fig. 19: (III)
Lateral view of a late Gt or early prophase nucleus with oriented chromosomes that
have rather uneven diameters and suggestions of coiling. Fig. 20: (III) Lateral view of
an early prophase nucleus with oriented chromosomes that have uneven diameters and
suggestions of coiling. Many of the telomeres are heterochromatic. Fig. 21: (III) Late
prophase nucleus with oriented chromosomes that have comparatively even diameters
and heterochromatic telomeres. Figs. 22—29. Nuclei that have been aceto-orcein
stained according to schedule I in Table 1. Fig. 22: Telophase nuclei in which chromosomes are still visible to some extent. Fig. 23: On the left is a late telophase or early Gj
nucleus in which chromosomes remain visible. On the right is a late Gx nucleus in which
chromosomes are not visible in the comparatively diffuse chromatin. Fig. 24: Labelled
early to middle S phase nucleus in which chromosomes are not visible in the diffuse
chromatin. Fig. 25: Labelled late S phase nucleus in which chromosomes are not
visible in the diffuse chromatin. Fig. 26: Early G, nucleus in which chromatin appears
in clumps and thin strands. Fig. 27: Late G% nucleus in which chromatin seems to be in
more distinct strands than in Fig. 26. Fig. 28: Early prophase nucleus in which the
chromatin is in the form of thick chromosomes with irregular diameters, and the
chromosomes have retained their relic anaphase-telophase orientation. Fig. 29:
Middle prophase in which the chromosomes are long and thin with more regular
diameters than in Fig. 28. Again the chromosomes can be seen to have retained their
relic anaphase-telophase orientation.
S. M. Stack, D. B. Brown and W. C. Dewey
290
47
48
49
50
Visualization ofinterphase chromosomes
291
of the same nuclei after aceto-orcein staining and after the sodium hydroxide treatment (schedule II in Table 1). Although boundaries between clumps of interphase
chromatin looked fuzzy after this treatment of A. cepa nuclei, the procedure worked
reasonably well for CHO cells. A comparison of Figs. 51 and 53 with Figs. 52 and 54,
respectively, indicates the sodium hydroxide treatment causes some swelling of
chromatin and the appearance of distinct chromatin domains that may correspond to
interphase chromosomes.
As a test of the hypothesis that the 10-30-nm chromosomal fibres must have
numerous attachment sites to the nuclear envelope in order for chromosomes to
reorganize at prophase (Comings, 1968, and see Discussion), A. cepa and CHO cells
were examined in prophase by both light and electron microscopy. As the hypothesis
predicts, prophase chromosomes generally are condensed against the nuclear envelope
in CHO cells (Fig. 55). However, in conflict with the hypothesis, A, cepa prophase
chromosomes condense throughout the nucleoplasm (Figs. 56-58). Apparently in
A. cepa nuclear envelope attachment sites are not necessarily needed for the reorganization of chromosomes during prophase.
Figs. 30—50. Chromosomes and nuclei from Chinese hamster cells (CHO line). The
nuclei in all figures except 47 were obtained from synchronized cultures. The
sampling time is given in parentheses at the beginning of the description of each figure,
x 1500. Figs. 30-47 illustrate sodium hydroxide-treated nuclei (schedule IV in Table 1).
Fig. 30 ( o h ) : Polar view of a metaphase chromosome spread. The arrangement
of chromosomes is characteristic of a hollow spindle. Fig. 31 (oh): Polar view of early
telophase chromosomes that have the arrangement characteristic of a hollow spindle.
Fig. 32 ( o h ) : Middle telophase in which the chromosomes begin to swell.
Fig- 33 ( o h ) : Late telophase or early Gx nucleus in which the chromosomes continue
to swell and uncoil (arrow). Fig. 34 (1 h): Middle Gj nucleus in which the chromosomes have swollen unevenly along their lengths (arrow). Fig. 35 (3 h): Late G1 nucleus
in which the chromosomes are less distinct than in Fig. 34. Fig. 36 (6 h): Early S-phase
nucleus in which much of the chromatin is rather diffuse, but a few chromosomes
are still in the process of uncoiling (arrows). Fig. 37: Autoradiograph of the nucleus
in Fig. 36. Fig. 38 (6 h): Early 5-phase nucleus in which the chromosomes are even
more loosely coiled (arrow), but many distinct clumps of chromatin remain. Fig. 39:
Autoradiograph of the nucleus in Fig. 38. Fig. 40 (6 h): Nucleus in middle S phase in
which part of the chromatin is finely divided while the rest is still clumped. Fig. 4 1 :
Autoradiograph of the nucleus in Fig. 40. Fig. 42 (1 o h): Late S-phase nucleus in which
thick chromatin strands begin to reappear. Fig. 43. Autoradiograph of the nucleus in
Fig. 42. Fig. 44 (12 h): Lateral view of a G a nucleus in which thick chromosome-like
structures are visible in their relic anaphase-telophase orientation (arrows). Fig. 45
(12 h): Mid- to late G, nucleus in which chromosomes have contracted into distinct
thickened strands with uneven diameters. Apparently contraction like swelling (see
Fig. 34) does not occur at the same rate along the length of chromosomes. Fig. 46
(12 h): Early prophase nucleus in which some of the chromosomes have contracted to
very thin strands, but many thick regions remain. Fig. 47 (non-synchronized): Prophase nucleus in which the chromosomes are thin and have comparatively even
diameters. Figs. 48-50: Aceto-orcein-stained nuclei photographed prior to the
application of autoradiographic emulsion. These nuclei have a fine network of
chromatin fibres with clumping more evident during Gx and G, compared to S phase.
Fig. 48 (1 h): Gj nucleus. Fig. 49 (10 h): Middle (left) and late (right) S phase nuclei.
Fig. 50 (12 h): G2 nucleus.
S. M, Stack, D. B. Brown and W. C. Dewey
58
Figs. 51-54. Nuclei from non-synchronized CHO cells prepared according to
schedule II in Table 1. Autoradiography was not performed on these nuclei, x 1200.
Fig. 51: Aceto-orcein-stained telophase (lower arrow) and Gx nuclei (upper arrows).
In the Gj nuclei the chromarin is in a fine reticulum while in the telophase nucleus thin
chromosomes are visible. Fig. 52: The same nuclei illustrated in Fig. 51 after sodium
hydroxide treatment. Apparently the chromatin domains in the G, nuclei and chromosomes in the telophase nucleus swelled after the sodium hydroxide treatment. Fig. 53 :
Aceto-orcein-stained S phase (left) and late G 2 or early prophase (right) nuclei. The
chromatin is in a fine reticulum in the S-phase nucleus, but thin chromosomes are
visible in the late G, or early prophase nucleus. Fig. 54: The same nuclei illustrated
in Fig. S3 after sodium hydroxide treatment. Some clumping of chromatin is visible
in the 5-phase nucleus and distinct strands are visible in the late G, or early prophase
nucleus. Again it is apparent that the sodium hydroxide treatment induced some
swelling of chromarin. Fig. 55: Electron micrograph of a CHO cell in prophase
showing that chromosomes condense primarily along the nuclear envelope, x 5000.
Fig. 56: Electron micrograph of A. cepa root rip cell in prophase showing condensation
of chromosomes throughout the nucleoplasm. x 7000. Figs. 57, 58: Light micrographs
of sectioned A. cepa root rip cells in prophase showing condensation of chromosomes
throughout the nucleoplasm. x 1200.
Visualization of interphase chromosomes
293
DISCUSSION
Concern about the physical state of chromosomes during interphase can be traced
back to the old arguments over whether chromosomes even exist as separate entities
during interphase. Although subsequently the persistence of chromosomes as units
has been demonstrated adequately by both genetic and morphological evidence
(Boveri, 1909; Wilson, 1928), the physical state and boundaries of chromosomes
during interphase remain unclear.
The most common means of visually examining chromosomes and nuclei at present
involve light microscopy of squash preparations and electron microscopy of thin
sections and whole mounts. Feulgen, aceto-orcein, and aceto-carmine-stained squashes
generally do not reveal structures in interphase nuclei of higher plants and animals
that are recogni2able as chromosomes. Thin sections and whole mounts of interphase
and non-mitotic nuclei examined in the electron microscope show a tangle of 10-30-nm
fibres attached at numerous sites to the nuclear envelope (Feldherr, 1972). Lafontaine
& Lord (1974) recorded changes in the pattern of clumped chromatin in A. porrum
root tip nuclei through interphase, but it is difficult to relate these changes to the
structure of individual chromosomes. In A. cepa and CHO cells chromatin fibres
become increasingly dispersed from Gx through S, and at least in A. cepa the
chromatin fibres condense somewhat during G2 (Dewey et al. 1972; de la Torre et al.
1975). In all, the results from electron microscopy of interphase nuclei have been
scanty and difficult to interpret, but in a recent review Wischnitzer (1973) comes to
the positive conclusion that the results of electron microscopy as early as the 1950s
' ...established that discrete interphase chromosomes are absent'. In agreement with
this interpretation, recent models for the structure of chromosomes during interphase
assume a general uncoiling of the 10-30-nm fibres of which chromosomes appear to
consist (DuPraw, 1970) so that the main pattern of chromosome 3-dimensional
structure is maintained only by numerous attachments of the fibres to localized regions
of the nuclear envelope (Comings, 1968; DuPraw, 1970; Brasch & Setterfield, 1974).
As the chromosomes begin to condense during prophase the fibres are somehow
retrieved from intermingling in the nucleoplasm to reform chromosomes against the
nuclear envelope (Comings & Okada, 1970a, b).
In apparent conflict with this interpretation of the structure of chromosomes during
interphase, there is an extensive literature based mainly on light microscopy of living
cells and cells that have been fixed and sectioned that suggests chromosomes do not
lose their 3-dimensional structure during interphase.
The chromosomes of some algae and protozoa remain distinctly visible through
interphase (DuPraw, 1970; Grell, 1973). However, because of the long phylogenetic
separation of these organisms from higher plants and animals and their possession
of certain other differences in mitosis that are considered anomalous or primitive,
these cases are not ordinarily used as evidence for the condition of interphase chromosomes in higher organisms. But there are special cases among higher plants and animals
in which chromosomes do remain visibly distinct during interphase.
Chromosomes are said to be visible in the spermatozoa of 2 nematodes (Meves,
s M Stack
294
- > D- B- Brown and W. C. Dewey
1915; Mulsow, 1912). Large polytene chromosomes of both animals and plants remain
distinct through endomitotic interphases (Brown & Bertke, 1974). Sex chromatin is
a well known example of the visibility of one heterochromatic X chromosome through
interphase in somatic cells of mammals (Barr & Bertram, 1949; Davidson & Smith,
1954), and constitutively heterochromatic parts of chromosomes generally are reported
to be visible through interphase in both higher plants and animals (Yunis & Yasmineh,
1972; Stack & Clarke, 1973). Finally, in some instances chromosomes may form
separate 'vesicles' during telophase, and these 'chromosomal vesicles' may remain
distinct but associated through interphase. Usually this has been reported in rapidly
dividing cells such as occur in cleaving animal embryos and spermatogonia (Wilson,
1928), but 'chromosomal vesicles' have also been reported to constitute interphase
nuclei in normal and malignant fibroblasts of the rat and mouse (Lewis, 1948), nonmitotic cells of frogs and rats (Kater, 1927,1928), and Phaseolus (bean) root tips (Kater,
1926). In these cases the authors usually assume that each chromosome is surrounded
by its own membranous envelope. This may be the case for some cleavage and possibly
spermatogonial cells, but it seems unlikely for the other examples (Ris & Mirsky,
1949). Still, regardless of the authors' explanations of how the chromosomes remain
visible, they claim to be able to observe chromosomes as individual units through
interphase.
Several different experimental treatments of living but primarily non-mitotic cells
of a variety of plants and animals have been reported to cause chromosome-like
structures to appear in nuclei. Effective treatments for specific cell types include
pricking nuclei with sharp instruments (Chambers, 1924; Ris & Mirsky, 1949),
treating cells with most of the common fixatives that contain acetic acid (Chambers,
1924; Ris & Mirsky, 1949), and exposing cells to solutions that vary in dissolved salts
(Anderson & Wilbur, 1952), pH (Philpot & Stanier, 1956), and tonicity (Kuwada &
Nakamura, 1941; Chambers & Black, 1941; Ris & Mirsky, 1949). These observations
were consistently interpreted to indicate that intact nucleoplasm consists of a gel of
hydrated and intimately associated chromosomes that can be dehydrated by various
means to reveal structures that are similar to prophase chromosomes. It has also been
suggested that such chromatin condensations may be comparable to the condensation
of chromatin during mitotic prophase (Chambers, 1924; Philpot & Stanier, 1956).
However, since most of these studies were on non-mitotic cells, the nuclei were probably structurally comparable to Gt interphase nuclei (Brown & Bertke, 1974), which
ordinarily do not condense into prophase chromosomes. One can still interpret this
work to indicate that chromosomes of some non-mitotic nuclei and possibly Gx nuclei
remain 3-dimensionally intact but swollen.
Support for this interpretation of chromosomes in interphase and non-mitotic cells
comes from a variety of other sources. Chambers & Black (1941) described the
chromatin in the normal non-mitotic nuclei of living onion bulb scales andTradescantia
leaf hairs as resembling, ' , . . . brain coral, with sinuous winding strands without beginning or end and filling the entire nucleus except for one or two rounded nucleoli'.
Interphase and non-mitotic chromosomes have been isolated from the nuclei of
a variety of vertebrate tissues (Claude & Potter, 1943; Mirsky & Ris, 1947, 1951;
Visualization of interphase chromosomes
295
Yasuzumi, 1951; Van Winkle, Renoll, Garvey & Prebus, 1952), leukaemic human
cells (Polli, 1953), and insect muscle and follicle cells (Pfeiffer, 1950). In these cases
unfixed nuclei were broken, and ehromatin in the form of recognizable chromosomes
was isolated. The chromosomes consisted of one or two chromatids that were longer
than normal metaphase chromosomes, and these chromosomes could be induced to
swell or contract depending on the medium in which they were suspended (Mirsky &
Osawa, 1961). Finally, our own observations of sodium-hydroxide-treated G± nuclei
also indicate the nucleoplasm consists largely of elongate, swollen chromosomes.
More recently, complementary evidence concerning the state of chromosomes
throughout interphase has come from experiments on fusion of mammalian nonmitotic or interphase cells to metaphase cells utilizing Sendai virus. Such fusion often
results in 'premature chromosome condensation (PCC)' in interphase nuclei (see
Sperling & Rao, 1974, for a review). In as little as 5-10 min after fusion the nuclear
envelopes of interphase cells break down, and the ehromatin in G1 and G2 nuclei
condenses to form recognizable chromosomes with one or two elongate chromatids,
respectively. S phase ehromatin does not condense appreciably, but apparently spills
out of nuclei as very long intact strands that probably correspond to individual
chromosomes (Rohme, 1975; Sperling & Rao, 1974). Fusion of metaphase cells to
non-mitotic cells such as lymphocytes, spermatozoa, and erythrocytes results in
Gj-type PCC (Johnson, Rao & Hughes, 1970). Similarly, G^type PCC is observed
in brain nuclei injected into Rana pipiens primary oocytes at the time of germinal
vesicle breakdown (Ziegler & Masui, 1973). The appearance of interphase chromosomes after sodium hydroxide treatment is similar to prematurely condensed chromosomes with the exceptions that prematurely condensed chromosomes are better
separated from one another and can be seen to consist of 2 chromatids at G2- These
2 differences are expected, since an intact nuclear envelope holds sodium hydroxidetreated interphase chromosomes together and, for reasons that are not clear, the
sodium hydroxide technique does not allow the visualization of sister chromatids
even at prophase or metaphase (Figs, i, 20, 21, 30, 46, 47).
If one assumes that all of these techniques for observing interphase chromosomes
are probably revealing comparable structures in interphase nuclei, an important
question is how the sodium hydroxide technique allows visualization of interphase
chromosomes, particularly when it does not employ pretreatment or even observation
of living nuclei as most of the other methods do. Although we have only begun to
examine this question experimentally, we suspect the successive treatments cause
shrinkage and swelling of ehromatin that results in its being visibly separated into
distinct domains or clumps that correspond to individual chromosomes. The use of
acetic acid in fixation has been reported to cause condensation of ehromatin during
interphase (Ris & Mirsky, 1949). Following squashing and air drying, the slides are
immersed in a dilute solution of sodium hydroxide that probably causes the ehromatin
to swell (Drets & Shaw, 1971; Comings, Avelino, Okada & Wyandt, 1973; and see
Figs. 51-54). The slides are quickly washed in 70 and 95 % ethanol which should
dehydrate the ehromatin and cause it to contract strongly (Willey, 1971). Finally
the brief incubation in 2 x SSC may allow a partial swelling and restoration of
296
S. M. Stack, D. B. Brown and W. C. Dewey
chromosome structure after the extremes of swelling and contraction produced by
the previous steps.
The preceding descriptions of metaphase nuclei also suggest an explanation for the
failure to observe interphase chromosomes as individual units by electron microscopy.
Unlike the sodium hydroxide technique, fixation for electron microscopy is usually
a careful attempt to alter the structure of cellular components as little as possible. If
one assumes the nucleus is composed essentially of swollen chromosomes between
which there is ordinarily no space other than that occupied by nucleoli, the 10-30-nm
chromosomal fibres may intermingle to some extent on the borders of chromosomes.
This would blur the boundaries of chromosomes at the levels of both light and electron
microscopy. Since most of the chromatin of the nucleus has the same density,
refractive index, and staining properties, it would be surprising if individual chromosomes could be seen without somehow separating them from one another.
With reference to present models for interphase chromosome structure, it seems
clear that complete decondensation of chromosomes does not occur in the intact
interphase nucleus, but rather the chromosomes swell while otherwise maintaining
their relic anaphase-telophase arrangement. Furthermore, attachment of chromosome
fibres to the nuclear envelope probably is not the sole basis of maintaining the structure
of chromosomes through interphase, because many of the chromosomes of A. cepa
condense at prophase without apparent association with the nuclear envelope. This
observation may be related to the model for nuclear-envelope-dependent chromosome
condensation (Comings & Okada, 1970a, b) by the recent suggestion of Comings &
Okada (1976) that chromosomes condense in association with a protein network
called the 'nuclear matrix' (Berezney & Coffey, 1976), that includes not only the
nuclear-envelope-pore complex but a fibrous network throughout the nucleoplasm.
The precise relationship between this fibrous network and individual chromosomes
is not yet clear.
Apart from its apparent capability of allowing some morphological study of interphase chromosomes, the sodium hydroxide technique may find further use in allowing
reasonable estimates of whether interphase nuclei are in Gv S or G2 without resorting
to tritiated thymidine labelling or microspectrophotometry. At present, apart from the
fluorescent technique of Moser, Muller & Robins (1975), there is no comparably
accurate and generally applicable technique based on staining (Alvarez & VaJladares,
1972; Dewse, 1974; Nescovic, 1968) or morphological differences (Nagl, 1970;
Lafontaine & Lord, 1974; Das & Alfert, 1968; de la Torre et al. 1975) to identify the
stages of interphase.
This research was supported in part by grants to S.M.S. from the Donald F. Jones Fund of
Research Corporation, and from the Faculty Improvement and Biomedical Support Committees
at Colorado State University and, to W . C D . from NIH Grant CA 08618.
We thank Dr T. C. Hsu and Dr Frances Arrighi in the Cell Biology Section of the University
of Texas M. D. Anderson Hospital and Tumor Institute at Houston, Texas, for warning us
of the problem in attempting autoradiography of Giemsa stained material, Dr Larry Hopwoodf
for assistance in autoradiography, Dr Marny Barrauf and Kathy Packard* for electron micrographs of CHO and A. cepa nuclei in prophase, and Dr David E. Comings in the Department
Visualization of interphase chromosomes
297
of Medical Genetics, City of Hope National Medical Center, Duarte, California, U.S.A., for
critically reading the manuscript.
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(Received 23 August 1976 - Revised 7 January 1977)