In vitro delayed senescence of extirpated buds from zooids of the colonial tunicate Botryllus schlosseri
Israel Oceanographic and Limnological Research, National Institute of Oceanography, Tel Shikmona, PO Box 8030, Haifa, Israel
* Author for correspondence (e-mail: buki{at}ocean.org.il)
Accepted 26 January 2004
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Summary |
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Key words: apoptosis, ascidian, Botryllus schlosseri, blastogenesis, budding, epithelial monolayer, life extension, senescence
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Introduction |
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The life history of a Botryllus schlosseri colony is
characterized, therefore, by multiple, blastogenic cycles, succeeding one
another periodically and each terminating in the takeover process.
Blastogenesis is intrinsically regulated and probably controlled by
blood-borne elements (Oka and Watanabe,
1957; Lauzon et al.,
1992
). Thus, Watkins
(1958
) demonstrated that
surgical removal of all buds from a colony did not affect the lifespan of its
functional zooids. Conversely, extirpation of all adult zooids in a colony
(Zaniolo et al., 1976
),
although resulting in a considerably belated development of many buds, did not
affect the lifespan and survival of its buds. Vascularization is an imperative
condition for this type of bud survival. When both functional zooids and buds
are removed simultaneously, vascular budding of replacement zooids may occur
(Milkman and Therrien, 1965
;
Sabbadin et al., 1975). Bud primordials then arise from the peripheral blood
vessels (the ampullae). This capacity is characteristic of many other
polystyelid ascidians (Rinkevich et al.,
1995
) and in some other colonial sytelids
(Watanabe and Newberry,
1976
).
Studies on blastogenesis have been performed at morphological levels
(Berrill,
1941a,b
,
1951
;
Oka and Watanabe, 1957
;
Watkins, 1958
;
Milkman, 1967
;
Izzard, 1973
; Sabbadin et al.,
1975; Zaniolo et al., 1976
;
Burighel and Schiavinato,
1984
), cytological ones (Lauzon et al.,
1992
,
1993
,
2002
), and
molecularbiochemical ones (Chang and
Lauzon, 1995
; Lauzon et al.,
1996
; Voskoboynik et al.,
2002
; Cima et al.,
2003
). These studies have revealed that blastogenic duration is
temperature-dependent and is shortened at elevated temperatures
(Milkman, 1967
; Sabbadin,
1969; Rinkevich and Shapira,
1998
; Rinkevich et al.,
1998
). Particularly severe conditions may involve regression of
adult zooids, irrespective of their blastogeneic stage
(Sabbadin, 1958
;
Rinkevich et al., 1996
). When
two ramets of the same genet or two allogeneically compatible genets at
different blastogenic phases are fused together, this difference is invariably
equalized (Watanabe, 1953
;
Rinkevich and Weissman, 1987
).
Ionization radiation (Rinkevich and
Weissman, 1990
) or acute administration of the anti-oxidant
butylated hydroxytoluene (Voskoboynik et
al., 2002
) may arrest blastogenesis, inducing a morphologically
chaotic state and deterioration that eventually lead to colony death.
Nevertheless, despite these several lines of intriguing observations, very
little is known about the mechanisms governing regulation of blastogenesis and
its control.
All the above studies were performed in situ or in vivo. In this study, we examined the in vitro fate of developing buds and regressing zooids isolated from B. schlosseri colonies at different blastogenic stages. Lifespan, apoptotic events, and cytological and morphological characteristics were compared with non-manipulated buds developing in vivo.
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Materials and methods |
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Primary buds at blastogenic stages `A' to `D' (sensu
Mukai and Watanabe, 1976) were
excised under the dissecting microscope by a sterile, pyrogen-free syringe,
equipped with a 28-G needle. Using the needle tip, we carefully peeled off the
uppermost layer of the tunic coating from around the buds without injuring the
buds' epidermis. During this manipulation, most secondary buds were
accidentally detached from the isolated primary buds. Buds from the same
colony were collected by sterile Pasteur pipettes and incubated for 30 min in
a 10 cm tissue culture Petri dish containing FSW-G at room temperature. The
buds were then transferred to a sterile nylon cell strainer (100 µm pore
size, Falcon, Becton Dickinson Labware, NJ, USA), immersed in FSW-G,
extensively rinsed with 200 ml FSW through the nylon filter and transferred
into a single 6-well culture dish (TPP, Switzerland) containing FSW
supplemented with 10% tunicate culture medium (TCM) for an overnight
incubation (20°C, 5% CO2 humidified incubator).
TCM was prepared as follows: 25 ml RPMI, HAM F12 or Fischer synthetic
liquid medium (single strength) was mixed with 19 ml double-strength
artificial seawater (ASW; stock solution 2 in
Rinkevich and Rabinowitz,
1993) and was supplemented with 4 mmol l1
L-glutamine, 20 mmol l1 Hepes, 3% heat
inactivated foetal calf serum (HI-FCS) and 1% mixed antibiotic solution
containing penicillin (10 000 U ml1), streptomycin (10 mg
ml1) and amphotericin B (25 µg ml1)
(PSA). A final volume of 50 ml was achieved by the addition of tissue culture
grade water. All media and medium supplements were purchased from Biological
Industries (Kibbutz Beit-HaEmek, Israel). All other chemicals were purchased
from Sigma (St Louis, MO, USA).
Buds, 26 per well, were then transferred into collagen/fibronectin pre-coated TPP 24-well plates or activated ProNectin 24-well dishes (Polymer Technologies Inc., San Diego, CA, USA), and incubated in a humidified incubator. An equal volume of 200 µl/well ASW supplemented with 3% HI-FCS, 2 mmol l1 glutamine and 1% PSA solution combined with 200 µl/well TCM, served as the initial growth medium. In the first 2 weeks, 50 µl of TCM were added to each well once every week, increasing the total volume to 500 µl/well. Thereafter, 50 µl of TCM/well were replaced once every 2 weeks, increasing nutritional concentration gradually.
Preparation of collagen and fibronectin-coated TPP-culture dishes
Type VII collagen from rat-tail (Sigma) was dissolved at a concentration of
1 mg ml1 in 0.1% acetic acid diluted with tissue culture
grade water. Collagen was allowed to dissolve overnight at 4°C. The
collagen solution was adjusted to a final concentration of 10 µg
ml1 by the addition of sterile tissue culture grade water. A
volume of 0.5 ml was pipetted to each of the 24-well culture plates,
vacuum-aspirated, air-dried and then polymerized in a laminar flow-hood under
UV light for 60 min. The plates were washed twice with sterile PBS and stored
at 4°C until use (up to 6 months). Prior to use, the collagen pre-coated
plates were incubated with medium containing fibronectin (5 µg
ml1) at 37°C for 3060 min. There was no further
addition of fibronectin whenever the medium was replaced.
MTT-colorimetric cell proliferation assay
Dissociated buds were incubated in 96-well plates with MTT-tetrazolium salt
(3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide; Sigma), at a
final concentration of 1 mg ml1 (4 h, 20°C, 175
µl/well). MTT salt was dissolved in TCM and then sterilized by filtration
(0.2 µm). The tetrazolium ring of MTT was reduced by active reductase
system (active only in viable cells) to water-insoluble formazan, blue in
color, that was subsequently solubilized by adding 100 µl/well extraction
buffer (total volume of 275 µl/well). Extraction buffer was prepared as
follows: 20% SDS was dissolved at 37°C in a solution of 50% dimethyl
formamide in DDW. The pH was adjusted to 4.7 by adding 2.5%, from 80% acetic
acid solution, and 2.5%, from 1 mol l1 HCl. Extraction
buffer releases the cell-bound dye when incubated overnight at 37°C.
Absorbance was read at 570 nm using an ELISA plate reader (TECAN, Spectra
Image, Austria). Background absorbance was established with wells containing
either extraction solvent and MTT without cells or extraction buffer with
cells only, omitting MTT. Five sets of experiments were performed and 200 buds
per experiment (at blastogenic stages `B' or `C') were isolated from subclones
of the same genotypes. Buds were enzymatically and mechanically dissociated,
generating approximately 104 cells per bud. Cells at a
concentration of 4x104 per well were cultured in 96-well
plates using two medium types: HAM F12 and Fischer.
DNA synthesis assay
Blastogenic stage `C' buds, isolated from the same colony, were grown in a
24-well collagen and fibronectin-coated dishes, three per well. To analyze DNA
synthesis, we exposed cultured whole buds to 5 µCi ml1
methyl-3H-thymidine (83 Ci mmol l1, 1 mCi
ml1; Nahal Sorek, Israel) (1 mCi=37 MBq) for 24 h at various
time points following bud extirpation. Buds were then repeatedly (x3)
rinsed with a total of 50 ml ASW/sample followed by a rinse of 20 ml ice-cold
10% trichloroacetic acid through a glassfibre profilter (25 mm, Sartorious, AG
W-3400, Goettingen, Germany). Each filter was then rinsed twice with 10 ml
anhydrous ethanol (20°C), air dried, placed in a separate vial and
3 ml per vial of liquid scintillation cocktail (Safe Fluor S Lumac/3M,
Netherlands) added. Tritiated thymidine incorporated into newly synthesized
DNA was measured by liquid scintillation counting.
PKH-26 assay for cell proliferation
Whole isolated buds were exposed to red fluorescent Zyn-linker PKH-26
(prepared for Sigma by Zynaxis, Inc.) as follows. Buds were rinsed twice with
1 ml of calcium and magnesium-free ASW in 15 ml polypropylene centrifuge tubes
(Greiner, Austria). PKH-26 solution was prepared at a concentration of
4x106 mol l1 diluted with Diluent-C,
which was provided with the kit. PKH-26-Diluent-C solution was added to the
buds, which were incubated for 10 min with gentle swirling to ensure a
homogeneous staining throughout bud tissues. Labeling was stopped by adding an
equal volume of 100% serum (foetal bovine serum, previously adjusted to
seawater osmolarity with NaCl), diluting the stain by half, and by adding an
equal volume of TCM. Finally buds were rinsed with TCM (x3, 10 min
each). All these procedures were carried out at room temperature. Thereafter,
the buds were cultured and observed under an inverted Olympus fluorescence
microscope equipped with an excitation filter of 550 nm and a barrier filter
of 590 nm. Zyn-Linker incorporated itself into cell membranes of the buds, and
remained stably incorporated for up to 2 months (C.R. and B.R., personal
observation). Proliferation is detected by the dilution of Zyn-Linker
fluorescence in daughter cells.
DNA fragmentation assay TUNEL, on attached monolayers
Isolated blastogenic stage `C' buds were cultured as described. The medium
from wells with developed epithelial monolayers was carefully aspirated under
a stereomicroscope and tissues were fixed in the well by adding 4%
paraformaldehyde solution (room temperature, 30 min). Paraformaldehyde was
first prepared at a concentration of 8% in DDW and then mixed with an equal
volume of 2x ASW. The fixative was replaced with 80% ethanol and the
plates were stored at 4°C until used. Staining was performed using the
Klenow-FragEL DNA Fragmentation Detection Kit (Oncogene Research Products, MA,
USA) basically following the manufacturer's procedures, with adjustments to
24-well plastic culture plates. Most solutions were added at a volume of 150
µl (the supplier recommends 100 µl). A minimum of 100 µl of the
Klenow Labeling Reaction Mixture was added (supplier recommends 60 µl) with
the aid of a coverslip. The coverslip was made of a round piece of parafilm
cut just slightly smaller in diameter than the well diameter. One edge of the
parafilm was folded up and was carefully applied on to the fixed tissue,
permitting the small volume of liquid in the well, to spread evenly.
Biotinylated nucleotides were detected using a streptavidin-horse radish
peroxidase conjugate. Dehydration was performed with absolute ethanol,
eliminating the step of xylene washes. A mounting medium was prepared by the
addition of nine volumes of glycerol to one volume of PBS; 200 µl of
mounting medium was added per well.
Histology of attached bud-bearing spheres
For histological observations and TUNEL staining, 1-month old in
vitro cultured buds (isolated at stage `C') with developed spheres were
fixed in tissue culture wells by 4% paraformaldehyde/artificial seawater for
30 min at room temperature. The paraformaldehyde was then replaced by 80%
ethanol. Specimens were subsequently dehydrated in a graded ethanol series and
embedded in paraplast. Sections 45 µm thick were cut with a rotary
microtome and stained either with Azan Heidenhain and Alum Hematoxylene and
Eosin or processed for TUNEL as described above.
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Results |
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In our in vitro protocols, buds did not attach to the substrates within the first 48 h after extirpation. But on day 3, approximately 50% of the buds isolated at stages `B' to `D' were adhered to the substrate. All buds isolated at stage `A' and some of the buds isolated at stages `B' to `D' remained unattached on the substrate. Both attached and unattached buds of stages `B' to `D' maintained their own heartbeat activity, which gradually ceased during the first week of explant cultures. With time, additional hemocytes migrated from lumens within the buds into the medium, appearing as aggregates or as single-cell suspensions (Fig. 2A). Elongated test cells with long pseudopodial extensions also migrated from explants (Fig. 2B), most abundantly from stage `D' buds, adhering tightly to the culture vessels. A few of them developed a dendritic morphology with several filamentous extensions. With time, small tissue fragments spontaneously dissociated from some explants. However, most buds maintained their external spherical integrity for the entire culture duration (up to 5 months).
|
Epithelium growth
Within 312 days of explant culture, we recorded growth of hollow
spheres (13 per bud; Fig.
2C) of a single epithelial layer in some isolated buds. Each
sphere was first noticed as a small vesicle that started to expand at an
average rate of 60 µm (diameter) per day, reaching up to 1000 µm (2
weeks). The spheres that remained unattached to the substrates deteriorated, a
process that developed within 50 days after they attained their full size.
Other spheres attached to substrates within 117 days of initial
appearance (attachment was also observed once in a 5 month-old culture) and
usually developed discoid monolayer of epithelial cells
(Fig. 2D). Monolayers grew
rapidly, primarily through cycles of cell division (PKH-26 stain, MTT assay
and thymidine incorporation, see below). The fast augmentation in cell numbers
initially formed confluent monolayer sheets (1520 µm cell size) that
expanded, especially at the periphery, into a monolayer of dilated (100 µm)
cells. Cells in the monolayers exhibited typical epithelial morphology, with
flattened polygonal lattice-like structures
(Fig. 2E). Peripheral cells
developed irregular structures, with long, thin cytoplasmic projections.
Epithelial sheets grew at an average rate of 230 µm diameter/day for up to
2 weeks and then ceased, starting a phase of cell deterioration. Cellular
deterioration was initially noticed by the appearance of a small number of
vacuoles around the nuclear membrane that expanded into the entire cytoplasm.
The attached intact monolayers began to dissociate within the next 3 days
leading to fragmentation of the sheet into small patches of cells
(Fig. 2F). Cells finally
rounded up and detached from the substrate.
Over a 2-month culture period, neither spheres nor attached epithelial monolayers developed from buds isolated at blastogenic stage `A' (5 experiments; 164 buds; Table 1). In isolated stage `D' buds (17 experiments; 201 buds), 36 of the 41 attached epithelial monolayers developed directly without going through a sphere stage. Approximately 2733% of buds isolated at stages `B' and `C' (N=430 and 429, respectively) developed spheres, of which only 8.6% and 6.5%, respectively, further developed attached epithelial monolayers (Table 1). These outcomes are also reflected by the ratios of monolayers to sphere, which are in an order of magnitude higher at stage `D' (5.12) than the 0.24 and 0.27 values at stages `C' and `B', respectively. In total, isolated stage `D' buds developed almost 3 times more epithelial monolayers than stages `B' and `C' buds. Surprisingly, some of the zooids isolated at mid takeover stage, just prior to parental heartbeat cessation, also developed epithelial monolayers within 820 days in culture (Fig. 3A,B; Table 2). These monolayers developed directly, as in most stage `D' isolated buds, without going through sphere development, and they were characterized by cells with larger nucleus-to-cytoplasm ratios (Fig. 3B) than in cells spreading from isolated buds. The deterioration of attached monolayers that developed from resorbing zooids (up to 10 days after development), and from isolated buds, is due to a nonapoptotic process, as revealed by the TUNEL assay (Fig. 3C). On the other hand, cells from other parts of cultured stage `D' zooids stained positive for apoptosis (Fig. 3E,F), including cells from the visceral tissues (pharynx, esophagus, stomach, intestine, endostyle), which had already been in an apoptotic state when isolated from the mother colonies.
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In cultures of stages `B' and `C' buds, spheres appeared at 312 days after isolation and survived for up to an additional 35 days. Attached epithelial monolayers appeared 325 days after bud isolation and survived for an additional 414 days (Table 2). Therefore, epithelial monolayers were present in cultures for up to 40 days after bud isolation, approximately a 1-month extension of their lifespan, compared to that of intact buds. This is equal to the duration of 45 blastogenic cycles. The attached bud's epithelium did not go through any massive apoptotic event (TUNEL; Fig. 3D). In some histological preparations of in vitro cultured buds with spheres, the majority of cells exhibited apoptotic nuclei; in other cases, they did not (Fig. 3GH). Internal morphology of these buds, both those with apoptotic and those with non-apoptotic cells, was distorted, exhibiting the loss of most visceral organ organization.
Cell proliferation
Exposing buds to PKH-26 resulted in a universal and uniform distribution of
dye binding to cell membranes. During cell divisions, the bound dye was
partitioned evenly between daughter cells, and so it gradually faded with
repeated divisions. In sphere tissues, cells lost fluorescent markers
(Fig. 4A,B), indicating cell
divisions there. Faint fluorescence was also observed at the base of each
sphere, at its point of contact with the buds
(Fig. 4B).
|
Cell proliferation was also documented by the MTT-assay in one of the five
experiments performed. In this experiment
(Table 3), epithelial growth
was observed in a single well at day 9 of culture and was characterized by a
significant increase (by a factor of 1.63) in enzymatic activity
(P<0.05; Duncan multiple range test), as compared to the other
wells at 1, 6 and 9 days. In the other four experiments, cells in cultures
exhibited low metabolic activities with average absorbance levels of 36. A 24
h exposure of buds to 3H-thymidine for 24 h at different
time-frames in vitro revealed low rates of incorporated thymidine in
cultures that did not develop spheres or attached epithelial monolayers,
indicating a low rate of cell division. After the first day in culture, the
isolated explant exhibited some proliferative activity
(Fig. 5); this was reduced
during prolonged in vitro conditions, as recorded previously
(Rinkevich and Rabinowitz
1993,
1997
). However, when sphere
development was observed, the cultures revealed a significant thymidine
incorporation (Fig. 5).
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Discussion |
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All previous studies (Oka and Watanabe,
1957; Watkins,
1958
; Milkman and Therrien,
1965
; Milkman,
1967
; Sabbadin et al., 1975;
Zaniolo et al., 1976
;
Rinkevich et al., 1995
;
Lauzon et al., 2002
) have
involved budectomized and/or zooidectomized assays, in which the developmental
responses of the regenerating colonies were observed and documented. The
present study reports the first experimental manipulations where buds and
regressing zooids have been surgically removed and followed under in
vitro conditions, away from any discrete colonial regulatory cue.
The application of this in vitro approach reveals several unexpected and intriguing results. (1) When spheres and epithelial monolayers appeared, the life expectancy of an isolated bud reached in vitro periods of 5060 days, exceeding by 5 times the life expectancy of intact, in vivo developing zooids. (2) The life expectancy of in vitro maintained buds that remained unattached to the substrates prolonged the above periods for up to 5 months. (3) After attaching to substrates, buds obeyed a newly developed `death clock', permitting up to 35 survival days for developing spheres and up to 14 days of life for epithelial monolayers. (4) The prevailing mode of death in vitro was necrotic, as opposed to the apoptotic mode of blastogenic takeover. (5) Isolated degenerating zooids under in vitro conditions produce epithelial monolayers within 3 weeks of culturing. Monolayers survived for up to 10 additional days, extending the lifespan of degenerating zooids from a few hours to 1 month.
In addition to the colony-wide weekly apoptotic cycles, B.
schlosseri colonies may also exhibit another type of colony-wide
programmed mortality, a non-random senescence
(Rinkevich et al., 1992;
Lauzon et al., 2000
). This
type of death is expressed cytologically as a series of necrotic events that
develop roughly simultaneously in multizooid clonal replicates experimentally
separated from colonies months earlier. We recently showed
(Voskoboynik et al., 2002
)
that the lifespan of the old, long-living phenotypes may further be extended
significantly by acute administration of the anti-oxidant butylated
hydroxytoluene.
The common vascular system (Burighel and
Brunetti, 1971; Zaniolo et
al., 1976
; Mukai et al.,
1978
) permits synchronization of colony-wide developmental
phenomena such as blastogenesis and non-random senescence
(Mukai, 1974
;
Burighel and Schiavinato, 1984
;
Lauzon et al., 1992
,
1996
;
Rinkevich et al., 1992
;
Voskoboynik et al., 2002
). The
determination of life or death is probably conveyed by blood-borne elements or
cues (Oka and Watanabe, 1957
;
Lauzon et al., 1992
;
Cima et al., 2003
) rather than
by the recycling macromolecular components from old to newly developed soma
(Lauzon et al., 2002
). In
experimentally manipulated colonies where different parts of the colony are
surgically removed, the profiles of colony-wide developmental responses
indicate a dynamic and synchronized central control for systemic changes
within the colonial boundaries. On the other hand, isolated buds or isolated
zooids at mid-apoptotic cycle are not liable to die as they are when
integrated into the colony. Under in vitro conditions, not only are
the underlying colonial mechanisms replaced by different developmental
entities (spheres, epithelial monolayers) and pathways, but also the colonial
level regulative longevity clocks are replaced by new biological clocks that
feature different developmental timetables (such as the 14-day
growth-to-senescence period for epithelial monolayers).
Studies that reveal alternative, non-apoptotic forms of programmed cell
deaths (e.g. Sperandio et al.,
2000; Leist and Jäätellä, 2001) or an array of
genes that significantly affect the lifespans of multicellular organisms
(Gems, 1999
;
Johnson et al., 1999
) may lead
to new insights into life-and-death programs and their roles in developmental
pathways. The in vitro investigation of Botryllus
developmental processes may provide an additional tool to understand clocks
that act on the whole-colony level as compared to life-and-death processes
expressed on the zooid level. These are two different tiers of developmental
signals that shape organ development differently in modular organisms.
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Acknowledgments |
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