historical perspective
Tracing the lineage of tracing cell
lineages
Claudio D. Stern and Scott E. Fraser
The study of cell lineages has been, and remains, of crucial importance in developmental biology. It
requires the identification of a cell or group of cells and of all of their descendants during embryonic development. Here, we provide a brief survey of how different techniques for achieving this have evolved over
the last 100 years.
he development of metazoan organisms is generally initiated by a series of
cell divisions that expand the number
of cells. In some species, even the earliest
divisions are accompanied by restrictions of
the developmental potential (their ability to
give rise to different tissue types) of the
newly generated cells. In other species, the
embryo just seems to increase its number of
cells, each of which remains multipotent
until a later stage when cell interactions
define their destiny. The former mechanism
(mosaic development) has also been called
the ‘European plan’1, because as in ancient
Royal houses, lineage ancestry is the primary determinant of the position of each
individual in society. The latter mechanism
(regulative development), in which the fate
of a cell is determined according to the
behaviour of its neighbours, has been
referred to as the ‘American plan’. It is now
becoming apparent that many organisms
use both mechanisms, but even for apparently similar processes, different organisms
can use different strategies. For example, in
amphibian embryos, early cleavage divisions subdivide the cytoplasm of the egg
into progressively smaller cells. During this
process, maternal ‘determinants’ of cell fate
are apportioned to different cells so that as
early as the third cleavage division, the cells
differ in the structures to which they can
contribute. By contrast, there is currently
no compelling evidence that early cleavage
divisions in mammalian embryos generate
cells with different potential; the cells of the
inner cell mass of the mouse embryo seem
to remain pluripotent for a considerable
time before becoming committed.
To understand how cells determine their
destinies in any developmental process, one
needs both to know the precise lineages of
cells in the normal embryo and to establish
the limits of the cells’ developmental potential (which can only be done by challenging
them, by placing them in new positions).
For both, it is important to have a reliable
method of identifying particular cells and
all of their descendants. It is tempting to
take advantage of ‘tissue-specific’ molecular
markers, but it has become increasingly
T
clear that very few markers are truly tissuespecific and, importantly, that cells of different origins can move in and out of
domains of gene expression, altering their
own expression pattern according to their
current location2. Because we cannot rely
solely on molecular markers, it is necessary
to define methods suitable for following
cells and their descendants, both during
normal development and after placing
them in new environments. This brief survey traces the history of the different
approaches for doing this.
The study of cell lineages can be said
with confidence to have started with the
seminal work of a small group of investigators (including Carl Otis Whitman,
Edmund B. Wilson and E.G. Conklin) at
a
b
the Marine Biological Laboratory in
Woods Hole, Massachusetts, at the beginning of the twentieth century. They studied early cleavages in embryos of different
invertebrates3,4. Conklin, for example, took
advantage of differences in pigmentation
between the early blastomeres of the tunicate embryo4. In theory, direct observation
of cells should be the preferred method for
studying cell lineages, because it does not
interfere with the developmental process
in any way. The advent of time-lapse cinematography (first applied with spectacular
results to living embryos by Wetzel5)
allowed the construction of lineage trees
by this method even when specific cells
could not be distinguished easily by their
morphology or pigmentation. The most
MB
FB
HB
OB
OT
Figure 1 Double-labelling with ‘DiO’ and ‘DiI’
illustrate that ablated regions and not the
unoperated neural folds form the neural
crest after ablation. a, Immediately after a
unilateral ablation of nearly half the neural
tube, the embryo received a focal injection of DiO (green) into the residual neural
tube in the midbrain and one of DiI (reddish orange) into the bordering neural-fold
cells in the hindbrain. b, After 36 h, the
neural tube cells (green) have dispersed
to form neural-crest cells in the midbrain.
In contrast, the existing neural-crest cells
(reddish orange) have migrated laterally
as expected for cells arising from rhombomere-4, entering the second branchial
arch. Only a few scattered cells deviated
rostrally from their original trajectory. The
optic cup (OP), forebrain (FB), midbrain
(MB), hindbrain (HB) and otic vesicle (OT)
are indicated for reference. (Modified
from ref. 40.)
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historical perspective
a
b
c
e
f
g
h
i
j
k
l
m
d
n
Figure 2 Fate-mapping the zebrafish neural
plate, illustrating the typical steps of a lineage-mapping study. a–d, First, a cell is
labelled with an indelible dye, and its position is carefully recorded in all three
dimensions. The accuracy of this information is essential for constructing prospective fate maps. e–m, The embryo is then
allowed to grow, and the positions of all
the descendants of the labelled cell(s) are
recorded. n, Finally, the results are put
together. The diagram on the left provides
a colour key for the identification of the
territories in the fate maps on the right:
red, telencephalon; black, diencephalon;
green, retina; orange, midbrain; blue,
hindbrain. The resulting fate maps show
the extent of the spatial domains at two
stages of labelling (upper diagram, 6 h
after fertilization; lower diagram, 10 h
after fertilization) that will eventually contribute cells to each of the identified
regions of the nervous system later in
development. By comparing the fate maps
obtained at these two stages, it is possible
to trace cell movements occurring in the
intervening 4 h. (Modified from ref. 41.)
dramatic example of the usefulness of this
method is represented by the tracing of the
complete lineage of the nematode
Caenorhabditis elegans, by direct observation and later by using ‘four-dimensional’
time-lapse video microscopy6,7.
The usefulness of the direct observation approach is obviously limited by the
ability of the observer to identify a single
cell and all of its descendants unambiguously at successive times, which is not
always possible, even with time-lapse cinematography. One common problem is that
in some cases it is not possible to see cells,
for example, when they have moved into
the interior of an embryo of an opaque
species. It was Walter Vogt8,9 who first
devised a systematic method for following
the fates of cells, using vital dyes (the word
‘vital’ implies that they can be used to stain
living cells, unlike most standard histological stains of his day). Vogt applied chips of
agar containing dyes of different colours to
the surface of urodele amphibian embryos;
small groups of cells would pick up the
dye, allowing their descendants to be identified even if at a later stage they were
located deep inside the embryo. A new
approach was introduced in the 1960s, taking advantage of radioactively labelled
compounds that had been introduced to
the marketplace. Small groups of cells
could taken from the embryo, labelled
with tritiated thymidine and then reintroduced into the embryo10.
One problem with vital dyes of the
types used by Vogt and Rosenquist is that
they are water-soluble, and it is therefore
difficult to exclude the possibility that they
could be transferred between adjacent but
unrelated cells. One solution was to construct inter-species chimaeras by transplantation, following the approach first
used by Spemann11,12 and Waddington13,
and later best exploited by Le Douarin and
her colleagues14,15. This method relied on
species-specific features that allowed
donor and host to be distinguished—by
pigmentation in Spemann’s experiments,
by cell size in Waddington’s, and by the
characteristic configuration of the
Feulgen-positive heterochromatin associated with the nucleolus of quail cells in Le
Douarin’s. Another solution resulted from
the introduction of carbocyanine dyes16,
which are lipid-soluble but water-insoluble and thus partition to the membranes of
cells. Two of these dyes, nicknamed DiI
(red) and DiO (green) quickly found
applications first for the tracing of axon
pathways17,18 and later for tracing cell lineages19–21 (Figs 1, 2).
Vital dyes, whether water-soluble or not,
are very easy to apply to cells. The experimenter needs only to bring them up to the
cells, which will take them up. But it is difficult to apply them to a single cell, and they
are therefore most often used to track the
fates of cell groups. To study the fates of a
single cell, the most direct method in the
absence of endogenous identifying features
is to inject an inert tracer directly into the
chosen cell. Two types of tracer have been
used extensively: the enzyme horseradish
peroxidase22,23 and fluorochromes conjugated to high-molecular-weight dextrans
(to prevent them from passing to adjacent
cells through gap junctions) and to lysine
(to make them fixable by aldehydes)24. This
method has been very successful for tracing
the fates of individual cells in vertebrates as
well as invertebrates, but it has the obvious
problem that injection into a single cell can
be technically challenging when the cell of
interest is small (nevertheless, the logic of
lineage-tracing experiments is most elegantly obvious when a single identified cell
can be labelled and all of its progeny identified). The additional disadvantage of using
enzymes or fluorescent compounds as tracers is that they become diluted with cell
division, which can become a severe problem in cells that divide rapidly or when the
study needs to be extended over a prolonged period of development.
To overcome both problems, an ingenious approach was devised25,26: a retroviral
vector is applied to introduce nucleic acid
encoding a marker (such as alkaline phosphatase or β-galactosidase) into cells. The
stock solution containing the viral particles is diluted so that infection events are
rare enough to maximize the probability
that all marked cells in a given embryo are
clonally related. To avoid the ‘secondary’
infection of cells not related to that initially marked, the retrovirus is made replication-deficient by the removal of sequences
coding for viral coat proteins. This
approach has been used successfully, particularly to trace cell lineages in the nervous system25, but it also has severe disadvantages. First, it only lends itself to ‘retrospective’ lineage mapping, because the cell
to be labelled cannot be chosen directly
(infection is a random event among cells
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historical perspective
expressing the appropriate receptor).
Second, a more serious problem is that
even when the viral stock is greatly diluted,
it is formally difficult to exclude the possibility that adjacent labelled cells are
derived from independent infection
events. To overcome this, Cepko and her
colleagues constructed a complex library
of about 80 different retroviruses 27.
Because infection is random, the chances
that two adjacent cells will be infected by
the same type of retrovirus by injection of
the library into tissue is thus greatly
reduced. Despite the much greater rigour
of this approach over conventional retrovirus-mediated lineage studies, the main
disadvantage remains that lineages can
only be analysed retrospectively, and additionally it requires that every cell group be
analysed by the polymerase chain reaction
to determine its relatedness to other
neighbouring labelled cell groups.
Another ingenious approach to tracing
cell lineages retrospectively uses a very infrequent, spontaneous DNA recombination
event to activate the expression of a marker
(β-galactosidase) in transgenic mice28. But
this recombination event occurs very early
in development, and the labelled cell can
therefore contribute to very many tissues,
complicating the analysis. To concentrate on
a particular tissue, such as muscle29 or the
nervous system30, a tissue-specific promoter
is added (so that the only cells detected are
those located within the chosen tissue that
are also descended from that in which the
recombination event occurred).
The methods for prospective lineage
analysis (vital dyes, transplantation and
intracellular injection of tracers) all select
the cells to be followed on the basis of their
position in the embryo at the time of
labelling. But it is sometimes useful to be
able to determine the fate of cells that
express a particular molecular marker,
which might not all be in the same place at
a particular time. One technique uses a
monoclonal antibody coupled to colloidal
gold to identify the descendants of cells
that express a common surface antigen,
regardless of their position in the embryo.
Antigen-expressing cells internalize the
antibody–gold complex and pass it on to
their descendants, where the marker can be
detected for at least a few divisions31. Later,
more sophisticated methods were introduced to achieve the same in ‘genetic’
organisms such as fly, nematode and
mouse. In mouse32,33, two transgenic lines
are generated: one carrying a marker gene
disrupted by a sequence delimited by LoxP
sites, and another line expressing Cre
recombinase under the control of a promoter that controls the expression of a
gene of interest. When the two lines are
crossed, the interrupted sequence between
the LoxP sites is excised, activating the
marker gene only in those cells in which
Cre is active, which corresponds to the
population of cells in which the chosen
promoter is active, irrespective of their
position in the embryo. In fly, this technique34 as well as the UAS-Gal4 system35
can be used.
This brief review has highlighted the fact
that there is no universally perfect method
for tracing cell lineages; the choice of the
best method must be dictated by the specific biological question being addressed.
Many developmental problems remain to
be studied that will require lineage analysis.
Although most of these techniques will
doubtless continue to be used, new methods continue to be developed. In addition
to elegant genetics, one promising
approach that is now being applied to many
species uses the electroporation of DNA
constructs into small groups of cells in
vivo36. Other exciting approaches include
the use of nuclear magnetic resonance
imaging to follow cells deep inside opaque
embryos37 and the use of spectral analysis to
discriminate between multiple dyes by twophoton microscopy38 (see http://bioimaging.caltech.edu/microscopy.html#spectroscopy). Increasingly, we are turning our
attention not just to tracing cell lineages,
but also to understanding how these lineages are coordinated with other dynamic
events such as cell movements and the spatial and temporal control of gene expression39. It will therefore be important to
develop methods of visualizing the behaviour of cells and their descendants at the
same time as the expression of particular
genes by the same cells and their neighbours. The next decade will undoubtedly
start to reveal some of the exquisite dynamics of cell behaviour within the living
embryo.
Claudio D. Stern is in the Department of Anatomy
and Developmental Biology, University College
London, Gower Street, London WC1E 6BT, UK
email: c.stern@ucl.ac.uk
Scott E. Fraser is at the Beckmann Institute (13974), California Institute of Technology, Pasadena,
California 91125, USA
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