Nucleoids and coated vesicles of “Epulopiscium” spp.

Arch Microbiol (1998) 170 : 227–235
© Springer-Verlag 1998
O R I G I N A L PA P E R
Carl Robinow · Esther R. Angert
Nucleoids and coated vesicles of “Epulopiscium” spp.
Received: 3 December 1997 / Accepted: 11 June 1998
Abstract We describe here aspects of the anatomy of
two “Epulopiscium” morphotypes, unusually large bacteria that are not yet cultured and that reproduce by the internal generation of two or more vegetative daughter cells.
Two morphotypes, A and B, which are enteric symbionts
of several species of herbivorous surgeonfish (Acanthuridae), were collected around the Great Barrier Reef of
Australia, preserved there, and later stained for light microscopy. Some samples were examined by electron microscopy. In both morphotypes, countless discrete nucleoplasms or nucleoids were found to occupy a single shallow layer just beneath the surface all around these organisms. At each end of the morphotype B cells, a membranebound compartment containing dense cords of chromatin
was observed. When these were found at each end of
growing daughter cells, no polar compartments were then
found in their mother organism. Electron micrographs of
sections of morphotype A symbionts show that their outermost region is composed of tightly packed coated vesicles, each surrounded by a thin, dense, spacious capsule.
Near the surface of type A organisms the remains of broken vesicles, broken capsules, and a finely fibrous matrix
fuse to form a fabric that serves as the cell wall. Morphotype B organisms, however, were observed to have a distinct, morphologically continuous outer wall.
Key words ”Epulopiscium” · Nucleoids · Polar
chromatin · Coated vesicles · Unusual composite wall ·
Daughter cells
C. Robinow (Y)
Department of Microbiology and Immunology,
Health Sciences Centre, University of Western Ontario,
London, Ontario, Canada N6A 5C1
e-mail:[email protected]
Tel. +1-519-661-3427; Fax+1-519-661-3499
E. R. Angert
Department of Molecular and Cellular Biology,
Harvard University, Cambridge, MA 02138, USA
e-mail: [email protected]
Tel.+1-617-495-0532; Fax+1-617-496-4642
Abbreviations DAPI 4′,6-Diamidino-2-phenylindole ·
PBS Phosphate buffered saline
Introduction
”Epulos” has been used as the name of members of a large
and varied group of uncommonly large bacteria (Clements
et al. 1989). The first such organism was reported from
the gut of the herbivorous surgeonfish Acanthurus nigrofuscus from the Red Sea. It was given the name “Epulopiscium fishelsoni” and was placed in the kingdom Protista
(Fishelson et al. 1985; Montgomery and Pollak 1988).
Various organisms sharing some of its properties were
later encountered in surgeonfish from Australia’s Great
Barrier Reef (Clements et al. 1989). Electron microscopy
of sections of the large organisms showed that they contain typically prokaryotic strands of chromatin and have
bacterial-type flagella. Their large size apart (some of
them attain a length of more than 500 µm), these bacteria
are unusual in that they multiply (or merely rejuvenate
themselves) in a viviparous mode of internal formation of
one, two, or more daughter cells that escape to the outside
through a tear in the wall of the mother organism (Montgomery and Pollak 1988).
Sequence comparisons of the 16S rRNA gene (Angert
et al. 1993) have placed “Epulopiscium” among the anaerobic, gram-positive, spore-bearing clostridia. More recently, Angert et al. (1996) have reported that the unusually large Metabacterium polyspora (Chatton and Pérard
1913), which is a symbiont of the cecum of guinea pigs
and other rodents and which forms two or more refractile
endospores per cell, is also located in this branch of the
genealogical tree. Spore-bearing cells of Metabacterium
spp. superficially resemble “Epulopiscium” morphotypes
engaged in the formation of vegetative daughter cells. Here
we describe observations made with the light microscope
on nucleoids and other forms of chromatin in “Epulopiscium” morphotypes. A brief account is also provided of
elements of the complex outermost layer or “cortex” of type
A organisms as seen in electron micrographs of sections.
228
Materials and methods
tail found to have been passably well-preserved, and this only in
the outermost regions of these symbionts.
Sample collection
Numerous samples of gut contents of surgeonfish were collected
on several occasions on Lizard Island (a part of the Great Barrier
Reef, Australia) by K.D. Clements. Samples were preserved there
in several different ways appropriate for 4′,6-diamidino-2-phenylindole(DAPI)-enabled fluorescence of DNA, the staining of
bacterial chromatin for light microscopy and electron microscopy
(see below).
DAPI fluorescence
Samples of gut contents were fixed and stored in 80% ethanol in
1993 and were recently prepared for the detection of DNA in the
following manner. Fixed samples were applied to poly-L-lysinecoated coverslips and air-dried. The coverslips were then immersed in a solution of 0.5 µg/ml DAPI in phosphate-buffered
saline (PBS) for approximately 2 min; subsequently, they were
rinsed with PBS. The coverslips were next mounted over a PBS/
50% glycerol solution. Slides were viewed and photographed using an Olympus BX 60 epifluorescence microscope as previously
described by Harry et al. (1995). Cells were visualized using an excitation cube unit (U-MNU) appropriate for viewing DAPI fluorescence with a narrow band-pass (360–370 nm) excitation filter and
a long band-pass (420 nm) barrier filter. Photographs of fluorescent preparations were obtained on Kodak T-MAX film, ASA 400.
Transmission light microscopy
The fixative for light microscopy of the large bacteria from surgeonfish was a mixture of two parts of a saturated aqueous solution of mercuric chloride with one part of absolute alcohol and
three parts water with acetic acid added to the mixture to make it
2% of the total volume. Fixation time was 10–15 min, after which
the sample was washed with and stored in 70% ethanol. Nucleoid
staining was achieved by both the Feulgen procedure and the HClGiemsa method (Piekarski 1937). Hydrolysis for 8–10 min with 1
M HCl at 60°C was followed by staining with the Giemsa mixture
of dyes. We used Gurr’s Giemsa “Improved 66” (British Drug
Houses, Pool, England) at one drop of stain per ml of distilled water that had been given a pH of 6.8 by the use of Gurr’s buffer
tablets (British Drug Houses). The progress of staining was
checked with a × 40 water immersion lens. Photographs were
taken of wet, stained preparations mounted over buffer of pH 6.8.
Electron microscopy
Samples intended for electron microscopy were prepared at the site
of collection in 2.5% glutaraldehyde/0.2 M cacodylate buffer (pH
7.2). Samples were fixed for 30–45 min, then washed with and
stored in 0.2 M cacodylate (pH 7.2) at 4°C. On arrival at the London laboratory, the deposit of centrifuged sample was gelled in
Noble agar and post-fixed for 1 h in 1% OsO4 in 0.2 M cacodylate
(pH 7.2), washed with water, and dehydrated via a series of
ethanol solutions of a concentration gradually increasing from 30
to 100%. Dehydrated samples in agar were next placed into Spurr
resin that was polymerized overnight at 60°C. Sections were cut on
a hand-operated Porter Blum microtome, were stained for 5 min
with uranyl acetate (2%) followed by 1 min in 1.4% lead citrate/1.8% sodium citrate, and were then examined in a Philips 300
electron microscope. The fixation procedures just described have
routinely provided satisfactory images of the membranes, nucleoids and ribosomes of bacteria studied in our laboratory, but the
interval between primary fixation of the giant symbionts in Australia and secondary fixation with osmium 5 weeks later proved to
have been too long. Only in sections of the A morphotype was de-
Results
The A morphotypes in our samples measured 200–250
µm in length, were slender, and tapered towards their tips.
Some of them bore one or two daughter cells, mostly at
the very early stages of development. B morphotypes
were only 100–185 µm long; they were straight cylinders
with rounded ends or were of slender cigar shape. Many
of these organisms bore one or two daughter cells.
Nucleoids of morphotype B
As Fig. 1 (A–E) and Fig. 2 (A,B) clearly show, B morphotypes have innumerable, closely-packed, small nucleoids
spaced all over their periphery in a shallow layer just beneath their surface. In DAPI preparations, brightly fluorescing polar compartments were invariably found at the
tips of free, mature individuals or in the tips of internally
arising offspring. These will be discussed below. At the
higher magnification of the Giemsa-stained preparation in
Fig. 3, we found that the nucleoid layer is composed of
large numbers of thin, flexible entities, nucleoids or nucleoplasms, packed together in single-layered, flat patches
of widely differing size and shape. Neighboring patches
are connected with strands or narrow ribbons also composed of nucleoids. Overall, the arrangement here is that
of a reticulate sheet – to borrow a term used by Smith
(1956) to describe the structure of the chloroplast of the
green alga Oedogonium. The nucleoids of B symbionts
vary in length and on the whole appear to be somewhat
smaller than those of common Bacillus species. After
Feulgen-style acid hydrolysis, the nucleoids of B morphotypes displayed an affinity for purple components of the
Giemsa stain, as in the case of other bacteria, but they did
so rather weakly. Oddly enough, they also proved to be
only weakly, if quite distinctly, Feulgen-positive in preparations in which the nuclei of protozoa, which were often
scattered among the giant symbionts, gave the expected
strongly positive response to the Feulgen procedure. In
retrospect, it seems possible that a shorter time spent in
acid hydrolysis would have evoked a stronger positive response of the B-type nucleoids to the Feulgen treatment,
particularly since DAPI scans showed the nucleoids of B
symbionts fluorescing as brightly as the nucleoids of other
bacteria contained in the same samples.
Densely chromatinic cords in membrane-enclosed space
at the tips of B-type symbionts
The weakly positive response of B morphotypes to the
Feulgen procedure is all the more remarkable because in
our samples, mature singles or pairs of daughter cells still
enclosed in their mother organism displayed, in the ma-
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Fig. 1A–E Epifluorescence of B morphotypes stained with 4′,6diamidino-2-phenylindole. A and B show median- and upper-surface optical sections, respectively, of the same cell. A The layer of
nucleoids just under the cell cortex is seen as a bright broken line.
DNA associated with putative daughter cell primordia is seen at
both poles. B A surface view of the network of islands of nucleoids
(A,B bars 50 µm). C and D show examples of B morphotypes with
mature daughter cells. Note the fine broken line of mother cell nucleoids surrounding the brighter daughter cell nucleoid layer. C
Two daughter cells in this image show coalesced DNA at the poles
and at regions along the side of each daughter cell. These may represent “granddaughter” primordia. D The unusual instance of a
single daughter cell. Coalesced DNA is seen at the poles of the
daughter cell but not in the mother cell. E A surface view of a lateral chromatin cluster of cords of DNA (C–E bars 20 µm)
jority of instances, at both poles small lens-shaped compartments containing several cords (which may be part of
a single knot) that were strongly Feulgen-positive and
also stained deeply after HCl-Giemsa (Fig. 4, A–C). We
shall refer to these structures as terminal or polar chromatin compartments. The cords of these compartments
are much larger and denser than individual nucleoids of
the superficial net shown in Fig. 2B. We believe that Fig.
5D of Montgomery and Pollak (1988) of a stout, twisted
cord of nucleoplasm in the center of what the authors regard as a developing daughter cell may represent part of
a knot of cords of chromatin within its compartment
similar to what we have observed at the tips of B morphotypes.
We can but speculate what may be the function of this
organelle that is generally present at both poles of the B
morphotype. Aware that daughter cells of some morphotypes of “Epulopiscium” arise in the tips of mother cells
[see, e.g., Fig. 6 of Montgomery and Pollak (1988)], we
propose that the terminal compartments with their prominent cords of chromatin may be primordia of daughter
cells. Among the DAPI-stained B-morphotype cells
scanned, many harbored long, slender daughters that invariably carried strongly fluorescing chromatin compartments at both ends, while no such compartments were
found in the remaining cytoplasmic space of their mother
organism even when, as in Fig. 1C, the mother cell’s own
superficial nucleoids were still vividly fluorescing. That is
what would be expected if, as suggested above, activated
terminal compartments transform themselves into daugh-
230
Fig. 2A, B Morphotype B (HCl-Giemsa). Views of the same
organism at two different levels of focus. A Median optical section. Note the straight cell wall of even density. The nucleoids appear as a line of pieces of chromatin just beneath the cell wall. The
dark, lens-shaped body at the tip is of a kind regularly found at
both poles of growing and of mature B morphotypes. Normally
these compartments contain stout cords of chromatin (see Fig. 4,
A–C). However, during the short interval between taking the photographs of A and B, the dark stain diffused away from the compartment and revealed in B – for reasons unknown – that the compartment was empty. B Overview of the superficial reticulate sheet
of nucleoids. The shreds of membranous materials clinging to this
cell are probably remains of the wall of its mother organism (bars
5 µm)
ter cells. For direct evidence of this proposed course of
events it would, however, be necessary to examine Btypes from much earlier phases of the growth cycle than
those represented in the samples that have been available
to us.
The DAPI preparations have also revealed instances of
cells that had, in addition to polar chromatin compartments, one or more brightly fluorescing clusters of chromatinic cords along their sides (Fig. 1, C and E). Such lateral clusters could perhaps account for certain B morphotypes mentioned by Clements et al. (1989) that harbor
more than two daughter cells. A look at 3-year-old Feulgen preparations in London revealed there, too, three instances of solidly Feulgen-positive, lateral chromatin
clusters among twenty B organisms. DAPI fluorescence
also drew our attention to the fact that even fully grown
type-B twin daughter cells often continue to cling together
for some time within the faintly illuminated remains of
the cell wall of their mother organism. Such remains are
Fig. 3 Morphotype B (HCl-Giemsa). Superficial net of clusters of
tightly packed nucleoids. The group indicated with an arrow is in
best focus (bar 5 µm)
seen clinging to much of the surface of the morphotype B
of Fig. 2B.
Nucleoids of morphotype A
In A morphotypes, the nucleoids were evenly distributed
over the entire subsurface plane of the cells (Fig. 5, A and
B). The nucleoids were not arranged there in any particular order, but were a remarkably uniform distance from
each other and quite possibly exist as distinct and separate
entities; differences in the size of individual nucleoids
may reflect their having been preserved at different stages
of their replication cycle. Near the tip of A symbionts, nucleoids tend to be more closely packed together than over
the rest of the body of these organisms, and some may extend from the plane of the main nuclear layer and enter
deeper regions of the cytoplasm. The nucleoids were
strongly Feulgen-positive, and most of them were slightly
larger than nucleoids of cells from fast growing cultures
of Bacillus mycoides and Bacillus megaterium [see Robinow
and Kellenberger (1994)].
Chromatin of internally generated daughter cells
The developing type-A daughter cells we have encountered were of two kinds. There were those whose chromatin presented as a tubular shell of seemingly solid chromatin perforated by large irregular holes (Fig. 5B). In the
231
Fig. 4A–C B morphotypes A, C HCl-Giemsa and B Feulgen.
Examples of polar compartments containing stout cords of chromatin (bars 5 µm)
other kind, which we regard as having been in a more advanced state of development, the chromatin was disposed
in much the same way as it was in the surface layer of the
mother cell except that the daughter’s chromatin tended to
stain more deeply than that of its mother organism. The
fluorescence of the chromatin of B morphotypes harboring juvenile cells seemed approximately equally bright in
mother organisms and offspring (Fig. 1C).
Light microscopy of the structure of the surface
of B and A morphotypes
Morphotypes B and A differ markedly in the nature of the
boundary between themselves and their environment. Optical sections such as that of Fig. 2A show that the type-B
symbiont is surrounded by a proper wall of geometrical
neatness and unvarying density. A narrow, translucent
zone intervenes between the wall and the nucleoid layer.
In contrast to this unexceptional pattern, we find the A
morphotypes of Fig. 5 (A and B) surrounded by a thick
“cortex”, to borrow a term adopted by Montgomery and
Pollak (1988) for the membranous outer boundary of “E.
fishelsoni” as a sign that they found it differed significantly from the cell walls of plants and bacteria. Note that
in our usage “cortex” will refer to all of the region exterior
to the layer of nucleoids. The type-A cortex of Fig. 5 (A
and B) seems to be of the nature of a soft rind that lacks
the sharply defined outer and inner contours of a proper
wall, varies slightly in thickness along the length of the
cell, and is not set off by a distinct gap from the nucleoid
layer. The complexity of this cortex was revealed by electron microscopy (Figs. 6, 7).
Fig. 5A, B Two A morphotypes (Feulgen). A Surface view of
shallow layer of evenly spaced nucleoids. Note along both edges
the rind-like, thick cortex. Out of focus in the interior is the dense
chromatin of a developing daughter cell. B Optical section of another A morphotype. In the polar region some nucleoids have entered deeper layers of the cytoplasm. Over the rest of the cell the
nucleoids are still seen as forming a shallow layer just beneath the
thick, translucent cortex. In the interior is seen the fenestrated tube
of dense chromatin of a developing daughter cell (bars 5 µm)
232
Fig. 6 Electron micrograph of
a section of the cortex of another A morphotype. The
cloud-like, faintly speckled
shapes near the bottom are profiles of nucleoids (n). Above
these is a labyrinth of interconnected spaces. Further up there
is yet another labyrinth where
the open spaces are lined with
much denser membranes (d)
than are those of the main
labyrinth below. The outer border of the cortex appears to be
formed by roughly circular
profiles of irregularly stacked
vesicles (bar 1.0 µm)
Electron microscopy of the cortex of A morphotypes
An overview of the periphery of a typical A morphotype
is provided in Fig. 6. Up from the micrograph’s lower
edge are seen profiles of nucleoids of low density and irregular contours similar to what has been found in E. coli
preserved only with gluteraldehyde, where, according to
Hobot et al. (1985), “… coarse aggregates of DNA were
present within a seemingly empty dispersed nucleoid.”
Continuing upwards, three levels of the outer cortex can
be seen. Prominent is an extensive layer of labyrinthine
design reminiscent of the maze of interconnected spaces
found by Clements and Bullivant (1991) just inside a
symbiont’s cell wall. Its upper reaches here are formed by
a shallow layer of open spaces whose lining membranes
are much denser than those of the main labyrinth below.
Higher still, a region of stacked ovoid or spherical structures is observed, and beyond these a layer of tightly
packed, ill-defined, perhaps fibrous materials are seen. Finally, exterior to this blurred horizon, a mat of bent and
broken filaments are observed; these have also been encountered at the periphery of “Epulopiscium” morphotypes studied by Clements and Bullivant (1991) and, on
the basis of their fine-structure, have been identified by
them as bacterial flagella.
Figure 7 shows a stretch of cortex similar to that of the
previous figure but at higher magnification. What is seen
in the higher levels of this micrograph and was found regularly in other sections of long stretches of uniformly
structured cortex of A morphotypes, we regard as profiles
of stacks of spherical or ovoid vesicles, each one enclosed
within a spacious, thin, dense membranous capsule.
Finely fibrous material appears to fill the space between
neighboring capsules. The vesicles within, with diameters
in the order of 0.11 µm, we regard as belonging to the
“decorated” or “bristle-coated” kind familiar to cell biologists. The closely packed decorated vesicles resemble the
“fluid segregating organelles” of Paramecium (McKanna
1976). These take the form of fascicles of narrow tubules,
blind at one end, that are situated proximal to the canals
233
Fig. 7 High magnification of a
short stretch of the outermost
region of the cortex shows
stacks of the characteristic profiles (thin arrows) of “decorated” or “bristle-coated” vesicles, each one surrounded by
the profile of its thin but dense,
spacious membranous capsule.
Below, thick arrowheads point
to vesicles preserved in the
process of being encapsulated
(bar 0.2 µm)
conducting fluid to the pulsating vacuoles. The constituent tubules of the fascicles bear an array of minute
pegs that endow cross-sections of them with a strong likeness to the bristle-coated vesicles of the outer cortex of
the A morphotype.
Discussion
Mode of reproduction of the A morphotype
The A morphotypes, like “E. fishelsoni” itself, reproduce
by the internal generation of daughter organisms (Fishelson et al. 1985; Montgomery and Pollak 1988; Clements
et al. 1989), but to date we have not found out where this
process initiates. Most probably the development of
daughter morphotypes starts at some point of the superficial nucleoid layer with its profusion of discrete nucleoplasms, but unambiguous evidence of such an event has
not been encountered in our samples. Among A morphotypes are found many that bear a single, short daughter
cell in the cytoplasm of the middle region of the mother
organism. Thus, a count performed on two Feulgenstained slides of A morphotypes from the same sample
yielded 27 that had single daughters varying from 28 to 40
µm in length and only 13 that bore two, long, overlapping
daughter cells. As to the origins of these pairs, there
would seem to be two possibilities. One scenario would
be that of two short daughter cells of the same length,
hence probably initiated at the same time (of which we
have seen examples), lying some distance apart, both
growing, eventually overlapping, and occupying most of
the available space. Another conceivable origin of overlapping pairs would be their having arisen from a single
short cell that had grown to great length and had, in the
end, divided by binary fission before extrusion from the
mother organism. To allow for the possibility of binary
fission of a single internally generated daughter cell
would not be unreasonable. It has, in fact, been observed
to take place in the cells of long, segmented filamentous
bacteria attached to the lining of the ileum of mice (Ferguson and Birch-Andersen 1979). Then again, two of
Montgomery and Pollak’s (1988) striking electron micrographs show a single, snake-like cell emerging from a torn
wall of the mother organism. If these scenes had been part
of a process of reproduction, then binary fission would,
presumably, in such instances have taken place soon after
extrusion. If not, what is observed here are instances of
mere rejuvenation and not propagation since the mother
organism is destroyed during the process. Uncertain is
also the life story, past and future, of the occasionally encountered instances of B morphotypes bearing a single
daughter within a mother organism whose own nucleoid
layer is still visible outside that daughter cell, as in Fig. 1D.
Coated vesicles of “E. fishelsoni”
The A morphotypes from surgeonfish around the Great
Barrier Reef (Australia), are not the only ones that are
equipped with coated or decorated vesicles. In electron
micrographs accompanying their ground-breaking paper
on “E. fishelsoni,” the giant enteric symbiont of surgeonfish of the Red Sea, Montgomery and Pollak (1988) describe “reticulate membranes and tubules at the cell periphery.” A close look at their work has convinced us that
the structures referred there to are what used to be called
“alveolate” vesicles or tubules. Some vesicles there have
234
a spiny circumference. Both aspects have been described
by Leedale (1967) in his writing about alveolate vesicles
near the pulsating vacuole of Euglena spirogyra. As he
put it: “Alveolate vesicles are distinctive structures, their
walls carrying a well defined alveolate patterning which
in sectioned vesicles appears as hairs radiating from the
surface”. We are aware that in current work in the field of
molecular cell biology, alveolated/reticulated vesicles
would be referred to as being “clathrin-coated,” but for
the time being we feel it appropriate to refer to such structures in the manner of the authors whose work we are here
discussing.
Coated vesicles are known to perform a variety of
functions not only in protists but also in the cells of certain tissues of plants and animals. Electron micrographs
do not reveal what coated vesicles contribute to the maintenance of the giant bacteria. Tentatively we propose that
their function is excretory, that in A morphotypes new
vesicles are steadily produced at the base of the vesicle
zone (Fig. 7) and are gradually pushed up to more peripheral levels and disintegrate there. The employment of the
bristle-coated kind of vesicles for the removal of waste
products may be a strategy that allows the A morphotype
to overcome obstacles that Koch (1996) sees as preventing bacteria from attaining large size. The continuous,
outward-moving supply of broken vesicles and fragments
of their membranous capsule, all seemingly embedded in
a fibrous matrix, may provide the cohesion required of a
layer that has to function as the equivalent of a cell wall.
Similar events seem to be happening in “E. fishelsoni.”
Montgomery and Pollak (1988) mention that the numerous tubules of this symbiont “frequently touch the inner
cortical surface” (i.e., the inner surface of the organism’s
bounding membrane), and so they do. But close scrutiny
of the relevant electron micrographs has provided clear
evidence that at several points the reticulated/alveolated
tubules actually fuse with and enter the substance of the
cortex and (perhaps only temporarily) impart to stretches
of it an alveolar pattern. Here, then, is another example of
vesicle/tubule-coating materials contributing something
to the building of a structure capable of performing the
functions of a wall. The course of events proposed here as
taking place near the surface of two kinds of enteric symbiont morphotypes may well seem improbable, but (in
morphological terms) it resembles the dynamics of the
steady loss of cell wall materials to the outside and their
renewal from below that has been demonstrated in grampositive, rod-shaped bacteria (Koch and Doyle 1985;
Beveridge and Kadurugamuwa 1996). We have not obtained unambiguous electron micrographs of B morphotypes, but have looked at enough of them to allow us to
say that the outer region of the cortex there is not composed of coated vesicles but of more sophisticated durable
structures and that B morphotypes have a distinct, twoply, coherent wall.
The samples of intestinal symbionts we examined were
from those segments of their life cycle through which they
pass during the hours of daylight. Thus, the few observations that we have been able to make leave much of the
life cycle of these giant symbionts unaccounted for. We
hope that others will collect over a stretch of 24 h a set of
anatomical/cytological observations on “E. fishelsoni” and
some of its morphotypes that would complement the circadian set of numerical data on growth and reproduction
collected by Montgomery and Pollak (1988). Meanwhile,
it is obvious that the nucleoids of the symbiont morphotypes we studied resemble those of common bacteria,
those of type A more obviously so than the strangely
small ones of the type-B symbionts. The validity of our
suggestion that the polar compartments containing stout
cords of chromatin may be primordia of daughter cells depends on the proof or disproof that can only be provided
by future life cycle studies. Until this information is available, we find support in the fact that, despite their large
size, these polar compartments resemble the passing stage
in the development of spores in Bacillus cells in which
distinct and variously coiled cords of chromatin are found
within a small compartment at the tip of rod-shaped cells
(Robinow 1960). Lastly, while the task of alveolate vesicles and tubules of “E. fishelsoni” and bristle-coated vesicles of the A-type cortex can at present only be guessed
at, it is of interest to have found that the employment of
such organelles is not limited to eukaryotes.
Acknowledgements The authors are much indebted to K. D.
Clements, now at the University of Auckland (New Zealand), for
catching surgeonfish and sending us samples for genetic analysis
and microscopy. For electron micrographs, the authors thank D.
Moyles, now at the University of Guelph (Ontario, Canada). C.
Robinow owes thanks to R. G. E. Murray for putting him on the
tracks of E. Angert’s work with the giant bacteria.
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