b-cell development in the amphibian xenopus
TRANSCRIPT
201
Immunological Reviews 2000Vol. 175: 201–213Printed in Denmark. All rights reserved
Copyright © Munksgaard 2000
Immunological ReviewsISSN 0105-2896
Louis Du PasquierJacques RobertMichèle CourtetRainer Mußmann
B-cell development in the amphibian Xenopus
Authors’ addresses
Louis Du Pasquier1, Jacques Robert2, Michèle Courtet1, Rainer Mußmann3,1Basel Institute for Immunology, Basel, Switzerland.2Department of Microbiology and Immunology, University of Rochester Medical Center, Rochester, New York, USA.3The Netherlands Cancer Institute, Division of Molecular Biology, Amsterdam, The Netherlands.
Correspondence to:
Louis Du PasquierBasel Institute for ImmunologyGrenzacherstrasse 487CH-4005 Basel SwitzerlandFax: 41 61 605 1222e-mail: [email protected]
Acknowledgements
We thank Dr M. F. Flajnik and Dr H. Etlinger for suggestions, comments and critical review of the manuscript. We thank Allison Dwileski and Lucy Trippmacher for the figures and the preparation of this manuscript. The Basel Institute for Immunology was founded and supported by Hoffman La Roche Ltd, Basel, Switzerland.
Summary: The amphibian Xenopus and mammals have similar organizationand usage of their immunoglobulin gene loci with combinatorial joiningof V, D and J elements. The differences in B-cell development betweenmammals and this amphibian are due to major differences in developmen-tal kinetics, cell number and lymphoid organ architecture. Unlike mam-mals, the immune system of Xenopus develops early under pressure todevelop quickly and to produce a heterogeneous repertoire before lym-phocyte numbers reach 5,000, thereby imposing a limitation on clonalamplification. In addition, it is submitted to metamorphosis. Thus, duringthe early antigen-independent period, several features of B-cell develop-ment related to immune diversification are under strict genetically prepro-gramed control: 1) D reading frames contribute complementary deter-mining region 3 with features that occur in mammals by somatic selec-tion, 2) the temporal stepwise utilization of VH genes in Xenopus occur infamilies probably because of structural DNA features rather than their posi-tion in the locus. Larval and adult immune responses differ in heterogene-ity. Larval rearrangements lack N diversity. During the course of immuneresponses, somatic mutants are generated at the same rate as in other ver-tebrates but are not optimally selected, probably due to the simpler orga-nization of the lymphoid organs, with neither lymph nodes nor germinalcenters resulting in poor affinity maturation. Switch from IgM to otherisotypes is mediated by loop-excision deletion of the IgM constant regiongene via switch regions which, unlike their mammalian counterpart, areA-T rich and reveal conserved microsites for the breakpoints.
Introduction
One of the best cold blooded vertebrate models for studying the
early development of the immune system is the (anuran)
amphibian Xenopus of the Pipidae family, also known as the
South African frog or the clawed toad (1–3). Since the last com-
mon ancestor to modern amphibians and mammals diverged
and became terrestrial at the end of the Devonian period 370
million years ago, Xenopus is a good model in deciding what is
accessory and what is essential in the immune system of verte-
brates. Indeed, what has been observed in both classes has to be
essential. As in mammals, the B-cell receptor diversity in this
species is generated via somatic rearrangement and combinato-
rial joining of multiple V, D and J elements within immunoglo-
bulin heavy and light chain loci (4). Xenopus development varies
202 Immunological Reviews 175/2000
Du Pasquier et al · B-cell development in Xenopus
partially from that of mammals and these variations appear to
influence B-cell differentiation as follows:
1) The tadpole immune system must develop its repertoire of
B and T-cell receptors (TCRs) as quickly and diversely as
possible since larvae hatch 2 days following fertilization. In
contrast to mammalian embryos, which remain in a rela-
tively antigen-free uterus, Xenopus larvae hatch in the sur-
rounding antigen-rich water.
2) When immunological competence occurs around day 12,
the number of cells is very small, with a few thousand B
cells and probably three times as many T cells. There is no
spleen prior to this stage and a response cannot be induced.
The differentiation events that take place in the immune
system depend either solely or essentially on intrinsic fac-
tors. This system is not in contact with the outside world
except, perhaps, via the blood, where there is no architec-
ture to guide the differentiation of cells. An equivalent stage
is reached in mammals when millions of lymphocytes are
already present.
3) There are few lymphoid organs both in Xenopus larvae and in
adults. The structures of these organs differ from those of
their mammalian counterparts and no germinal centers
have been described. Even in the adult, the total number of
lymphocytes differs in magnitude from that in mice. A
100 g Xenopus has approximately 1×108 lymphocytes (T+B),
whereas a 25 g mouse has twice this number in the spleen
alone.
4) The transition at metamorphosis when new adult-specific
antigens are expressed in a larval environment that is
immunologically competent is accompanied by many vari-
ations in lymphocyte phenotypes and function. This is
another natural experiment on the genesis of self tolerance
that does not exist in the same form in mammals. At this
time, compared with larvae, variations are seen in T–B col-
laboration in alloresponses and in antibody repertoire.
Because of these variations, studying the onset and evolution of
B-cell populations with or without specific immunizations in
Xenopus could assist in identifying essential versus accessory fac-
tors in the development of B cells in mammals.
The players – immunoglobulin genes, B cells and lymphoid
organs – in Xenopus
The Ig loci of Xenopus
Heavy chain – constant region
There are three isotypes of immunoglobulin heavy chain con-
stant region genes: IgM (4–6); IgY, a homolog of IgG (7, 8); and
IgX, preferentially expressed in the gut (9, 10). Their genes seg-
regate together but their order on the chromosome has yet to
be determined. The transmembrane and cytoplasmic domains
of Xenopus membrane-bound IgM (mIgM) are similar to the cor-
responding domains of all known vertebrate mIgM molecules.
The membrane forms of the two other Ig isotypes, mIgX and
mIgY, exhibit the specific structure found in all Ig membrane
exons but are not directly homologous with any specific mam-
malian non-µ Ig isotype; they are most similar to Xenopus mIgM.
Based on the conserved transmembrane domains of Xenopus
mIgX and mIgY, we believe that the heavy chain genes of IgM
and IgX initially arose from the original µ gene by means of
duplication. The transmembrane and cytoplasmic domains of
Xenopus mIgY have some conserved residues found in avian
mIgY and mammalian mIgG and mIgE, suggesting that the
more modern isotypes share a common but ancient ancestor
with amphibian mIgY (8).
Heavy chain – variable regions
Xenopus has approximately 100 VH genes belonging to 11 fami-
lies, more than 10 DH genes and eight to nine JH genes depend-
ing on the species (laevis: eight, gilli: nine) (11–13). The VH
locus is diploidized in the pseudotetraploid X. laevis as shown by
segregation studies (10) and by in situ hybridization with cDNA
probes and tyramide enhancing technology (Renaissance Tyra-
mide Signal Amplification, NEN Life Science Products) (Fig. 1)
(14). The VH genes are interspersed in the region (15) but not
scattered totally at random (4). A partial deletion mapping (L.
Du Pasquier, unpublished) using tumor cell lines that have
undergone different rearrangements of their Ig heavy chain loci
(16) showed that the VH8 and 5 genes are on the outside of the
locus, and a group encompassing most of the VH, i.e. VH1, 2 and
3 (at least 60 genes), is not close to the most 5' DH gene. The
sequence of the whole DH region is incomplete, but two
genomic D elements have been cloned (13). The distance
between the last D and the first JH is greater than 6 kb, the
amount of sequence upstream of JH determined without find-
ing a D element. The DH locus of Xenopus is therefore much fur-
ther upstream of the VH locus than in mammals. This could
explain some differences in the Ig rearrangement profiles of
Xenopus, such as the low incidence of DJ rearrangements (13).
Indeed, Yu et al. argue (17) that the distance between the seg-
ments can affect the kinetics of rearrangements of the different
elements, which could explain the observations in Xenopus.
Light chain
There are three isotypes of light chain, called �, � and �-like,
corresponding to products ranging from 25,000 to 29,000 Da
(18–21). The light chain locus architecture, numbers, and the
genetic linkages of the various loci are not determined with
Immunological Reviews 175/2000 203
Du Pasquier et al · B-cell development in Xenopus
precision except for the �-like r isotype. The latter’s architec-
ture is similar to that of mammals containing multiple V, at least
four Js and one C gene or segments (21). Compared with those
of � and �, the structure of the � J is different with the diglycine
bulge being replaced by a diserine (20).
B cells
These days, no cell exists that cannot be detected by flow
cytometry. Markers for Xenopus B cells are unfortunately scarce
except for heavy and light chain Ig (22). CD5, a marker for a
subpopulation of B cells in mammals (23), is a marker for T
cells in Xenopus but it is also detected on activated Xenopus B cells
after stimulation with phorbol 12-myristate 13-acetate (24).
So far, three stages: 1) pre-B cells (expressing heavy chain but
no light chain), 2) B cells, and 3) plasma cells have been chara-
terized in the spleen. The pre-B and plasma cells are of low den-
sity and heterogeneous in size while small sIg+ B lymphocytes
are of high density and much more homogeneous in size (25).
CD45, the leukocyte common antigen, is a structurally heter-
ogenous molecule ranging in molecular weight from 180 to
220 kDa. Xenopus B cells express the 220 kDa variant (26). Allelic
exclusion is the rule in Xenopus B cells, regardless of the number
of Ig loci present, following gene duplication by allopoly-
ploidization (27, 28).
Larval B cells differ from adult B cells in the re-expression
of surface Ig after capping. Whereas all adult liver and splenic B
cells re-express their Ig receptor 96 h after capping, only 30%
of larval B cells do so. Perhaps the cells that do not re-express are
immature B cells. After receiving a negative signal from anti-Ig
antibody these cells die as described for B cells from fetal mice
(29). Between larvae and adults the compartmentalization of B
cells changes. Adult immature B cells could perhaps be found in
adult bone marrow instead of larval spleen and liver (30).
Lymphoid organs of Xenopus
The spleen, which is also hematopoietically active in adults, is
the main site of B-cell differentiation (Fig. 2). The architecture
of the spleen shows a clear thymus-dependent area, with cells
labeled with anti-CD8 reagents (3, 31, 32) surrounding the
B-cell area around the central arteriola.
In accordance with data published so far in Xenopus, and in
general in cold blooded vertebrates (33), no structural equiva-
Fig. 1. Localization of the heavy chain locus by in situ hybridization. The chromosome preparation from the tumor cell line B3 B7 was hybridized with a cDNA probe from a VH1–IgM Xenopus clone labeled with digoxigenin (Dig) and revealed with anti-Dig antibody and enhanced by the tyramide method (L. Du Pasquier, M. Courtet, in preparation). Objective: ×100
Fig. 2. Immunohistochemistry microscopy on frozen spleen section at low magnification from an unimmunized Xenopus juvenile LG-15 double stained for IgM+ (blue) and BrdU+ (brown) cells. The frog was treated overnight by adding 2 mg/ml of 5-bromo-2' deoxyuridine (BrdU) in the water in darkness. Spleen sections were then incubated with anti-IgM 10A9 mAb followed by alkaline-conjugated goat anti-mouse secondary Ab preadsorbed twice on Xenopus blood cells. Following staining with AK substrate, BrdU incorporated in DNA was detected after DNA hydrolysis with hydrochloric acid using anti-BrdU mAb, horseradish peroxidase-conjugated goat anti-mouse secondary Ab and dimethyl-aminoazobenzene as substrate (brown cells). Notice the white pulp nodule in the center with the B-cell areas, stained in blue around the central arteriola, unlike those found in mammals. Objective: ×10
204 Immunological Reviews 175/2000
Du Pasquier et al · B-cell development in Xenopus
lent to germinal centers has been observed in the white pulp of
the spleen. Following immunization, specific antigen-reactive
cells were seldom found in clusters and were never associated
with structures resembling germinal centers (Fig. 2). Antibody-
producing cells were detected in the spleen as plasma cells but
no cluster of proliferating antigen-specific B cells was identified
by immunochemistry using 5-bromo-2' deoxyuridine (BrdU)
incorporation as an indicator of DNA synthesis (Fig. 3).
B cells are also in the adult gut (10) and thymus (34). In
contrast, the larval gut does not contain any B cells (i.e. Ig sur-
face positive). Even though compartmentalization is not very
pronounced in Xenopus lymphoid organs, thymus B cells seem to
switch more readily to IgY production upon T-cell help than
splenic B cells (34).
There are no lymph nodes in anuran amphibians, although
some lymphoid accumulations are found near the heart (33).
Rearrangement of Ig genes occurs in the liver and presumably
also in the spleen of tadpoles. Bone marrow is absent in tad-
poles. It was thought not to be lymphopoietic in adults (25)
but recombination-activating gene (RAG) activity (RAG trans-
cription) has been detected in it (35). Ig isotype production is
organ dependent, IgM is everywhere while IgG is not produced
in the gut. IgX, in contrast, is preferentially expressed in the gut
in intraepithelial plasma cells (10).
The lack of germinal centers typical of cold blooded verte-
brate lymphoid organs is thought to be responsible for the poor
affinity maturation in Xenopus (see somatic mutation section)
and could be due to the absence of classical follicular dendritic
cells (FDC). In mice, in the absence of FDC, germinal centers
are not formed normally, while affinity maturation, although
delayed, still occurs (36). Large thymus-independent dendritic
cells in the periphery of the splenic white pulp could represent
the Xenopus equivalent of FDC (37).
Further research in immune response regulation is now
required in Xenopus. Such research should involve the study of
antigen-presenting cells (FDC) and chemokines and their recep-
Fig. 3. A. Immunofluorescence microscopy on frozen sections of juvenile Xenopus LG-15 spleen. Detection of splenic B cells producing anti-2,4-dinitrophenol (anti-DNP) antibody, 3 weeks after immunization with DNP–bovine serum albumin (10 µg i.p.). Frozen section was incubated with DNP conjugated to alkaline phosphatase and stained with naphthol-AS-phosphate + fast blue as AK substrate. B. Spleen seen at low magnification (100×) with three white pulp nodules. Arrows point to putative antigen-specific plasma-cell like cells. C. Higher magnification (400×) of antigen reactive lymphocytes.
Immunological Reviews 175/2000 205
Du Pasquier et al · B-cell development in Xenopus
tors (38) involved in establishing the architecture of lymphoid
organs and the migration of helper cells. The same applies to
interleukins for which studies have just begun (39, 40).
Ontogeny – putting everything together
B-cell appearance and expansion during ontogeny
Xenopus larvae hatch 2 days after fertilization. From this stage
onwards, the young are exposed to the outside world and are
under pressure to develop a diverse functional immune system
rapidly. The liver and the thymus are colonized during the first
week of life by lymphopoietic precursor cells migrating from
the lateral plate mesoderm and ventral blood island. Lym-
phopoiesis occurring in late larval life and after metamorphosis
produces a stable, persistent population of ventral blood island-
derived stem cells as well as dorsally derived stem cells. Larval
stem cells can give rise to adult cell populations as shown by
ploidy marker experiments (41, 42). In Xenopus, developmental
studies of B cells are limited by the lack of markers. The pheno-
type of what is known as a B cell is always linked to the pre-
existence of a rearrangement, since only cells that have either a
heavy chain alone or a complete HL dimer on the surface can
be identified.
The development of the larval immune system in general
and that of B cells in particular can be divided into two periods:
one starts on day 4 when the first RAG message is detected and
ends on day 12 when the spleen can be identified. The second
period extends from day 12 to the end of metamorphosis,
approximately 50 days later. Before day 12, when the animal is
not yet immunologically competent, B cells are likely to
develop exclusively in the liver under the influence of intrinsic
determinants. Since light chain can only be detected from day
10 onwards, it is certain that each development and diversifica-
tion detected at the H chain level in the cytoplasm, prior to
then, is independent of external antigens. The spleen becomes
visible on day 12 and antibody responses can be elicited from
this day on (43). Extrinsic agents can now contribute to shap-
ing the immune repertoire.
Four to five days after fertilization half a dozen IgH chains
expressing immature B-cell precursors are detectable in the
liver of the larvae (Fig. 4). The number of immature B cells dou-
bles approximately once a day. After 2 weeks a few thousand B
cells are in the liver and spleen. At this stage the B-cell reper-
toire is very diverse, and each cell may express a different B-cell
receptor (44). This diversity presumably optimizes the ability
of young larvae to produce as many antibodies as possible
against a wide range of different antigens. This is consistent
with observations in another species of amphibian, the mid-
wife toad Alytes obstetricans, where during this developmental
period, cells specific to an early epitope are progressively
diluted by cells expressing different specificities (45).
During metamorphosis the lymphoid organs undergo a
drastic reduction in size and cell number and at the same time
RAG enzyme activity declines (46). A second histogenesis of
the lymphoid organ takes place during the month and a half
that follows metamorphosis and is most visible in the thymus
but to a certain extent also in the spleen (47). RAG enzymes are
active in adult, where they are detectable in the spleen and in
the bone marrow (35).
Rearrangement throughout ontogeny
RAG1 message is detected 4 days after fertilization at 22°C.
Heavy chain genes rearrange first, starting on day 5, followed
by light chains on day 8 (44). Until day 8 the developing
immune system is not functional because most IgM+ cells are
pre-B cells (48), with no L chain rearrangement having been
observed. VH genes then rearrange in a somewhat stepwise
manner (Fig. 4) (44). For L chain, only two loci out of three, �
and �� were precisely monitored during ontogeny, and � rear-
ranges earlier (18, 20, 44). If what has been observed in
tumors also occurs in normal B cells, rearrangement of � seems
to occur, as in mammals, with the deletion of � (16). No data
exists concerning a putative surrogate L chain in amphibians.
The stepwise rearrangements of H and L chain genes in
Xenopus (44) are different from those reported in mammals.
There is no positional effect. Different members of the VH1 fam-
ily can be used first and they are dispersed throughout the locus
(see previous section). The preferential expression of con-
served VH genes in different species must therefore be due to
common regulatory elements, for instance, recombination sig-
nal sequences. In Xenopus the sequence of events is compressed
over 5–6 days even though it retains the basic features found in
mammals. Those features are likely to be due to intrinsic factors
linked to the mechanisms of rearrangement. Indeed, the spacer
in the recombination signal sequences shows some family
traits, some of them conserved between Xenopus and mammals,
and they have been shown to be important in orienting rear-
rangement (49).
The Xenopus VH1 is homologous to the VH3 clan that rear-
range early in mouse and humans (50–53). However, the con-
served regions are within framework 1 and 3 and not within
the antigen-binding sites. This suggests that these gene seg-
ments are not selected on the basis of a common antigenic
specificity unless the antibodies using such VH bind antigens in
a different manner, for instance via the charge of the amino
acids in the � sheets rather than via the complementarity deter-
206 Immunological Reviews 175/2000
Du Pasquier et al · B-cell development in Xenopus
mining region (CDR) loops. In addition, VH1 rearrangement in
Xenopus occurs when L chains are not produced. Therefore, the
selective advantage of VH1 is unclear. In mammals, several mod-
els have been proposed to account for the usage of certain VH
during early ontogeny. For example, it has been reported that
fetal human and mouse B-cell repertoires are enriched for mul-
tireactive antibodies that exhibit low affinity binding to self
antigens (e.g. DNA antigens) (54). With the strict developmen-
tal control of the fetal and neonatal Ig repertoire, this led to the
hypothesis that the early generation of self-reactive antibodies
represents a critical feature of mammalian immune repertoire
development. A low level of self reactivity may serve as an
essential stimulant for B-cell selection and proliferation. Alter-
natively, the immune system developmental program, less strict
in Xenopus than in mammals, may reflect evolutionary pressure
to express a plastic set of antibodies, enabling an immunologi-
cally naive individual to respond to a wide range of antigens
but with a low affinity.
Very few D–J rearrangements (<1%) on the abortively rear-
ranged allele were isolated in adult and larva compared with
mammals, either at the cDNA or genomic DNA level (13). This
observation, which could be due to the position of the different
V, D and J loci (see above), does not leave much possibility for
the generation of a Dµ (55) protein in amphibians.
Ig heterogeneity throughout ontogeny
Heterogeneity of the rearrangement has been assessed from day
5 to adulthood (44). In isogenic Xenopus, all VDJ genes contain-
ing VH1 isolated from 5 to 7 day old LG15 larvae (when they
had 5–50 IgH chain but no IgL chain-positive cells) were dif-
ferent, showing the absence of clonal amplification. Diversifi-
cation appears to be maximized at this stage. At the genomic
DNA and cDNA level, the combinatorial and junctional diver-
sity of VDJ genes of 9–10 day old larvae were similar. The rela-
tively high frequency of non-functional out of frame rear-
rangements is typical of a developing population when precur-
Fig. 4. B cells during Xenopus development. Immunofluorescent staining with anti-� (open circles) and anti-L chains (black circles) has been lined up with the sequential rearrangements of VH (µ) and VL (� and �) segments. [Data from (44)]
Immunological Reviews 175/2000 207
Du Pasquier et al · B-cell development in Xenopus
sor B cells have not rearranged both Ig alleles and where the
ratio of functional to non-functional rearrangements reflects
the three possible reading frames (one in frame and two out of
frame). At a later stage (day 13–15), surface Ig+ B cells accumu-
late in lymphoid organs. Since selected B cells have now rear-
ranged both alleles, the ratio of functional to non-functional
rearranged VDJ genes in surface Ig+ B cells is now 1:1. Around
this period, rearrangements of all VH gene families (except
VH11, which is also rarely used in adults) were found. JH6 and
DH12 and 16 were observed for the first time in rearranged VDJ
genes. Finally, 38 cDNA sequences were obtained from tadpoles
at stages 56–58 (day 30–55). All were different (13). At the
same stage, nine different genomic rearrangements were iso-
lated from a B-cell genomic DNA library.
The diversity of sequences does not clearly account for the
restriction in antibody populations characterized by isoelectric
focusing (56, 57). Perhaps there is a microheterogenity in Xeno-
pus antibody sequences not necessarily linked to a charge differ-
ence which results in an apparently restricted response (58).
Even in the absence of N diversity, the CDR3s of developing
Xenopus are quite diverse at the cDNA level, but it is difficult to
correlate this diversity with that of antibody. Plasma cells pro-
duce an enormous amount of antibody that can mask the het-
erogeneity present in B cells at the protein level.
Secondary rearrangements
Secondary rearrangements leading to V gene replacements have
been found in Xenopus L chain genes within a circular DNA library
(59), although no functional studies have been performed to see
if this results in a selective advantage. In species with low cell
numbers, the selective value of this mechanism appears to be
evident, while in species with ongoing rearrangements the util-
ity is less obvious. The economical argument, that receptor edit-
ing saves B cells, could apply to frogs at metamorphosis, where
there are fewer B cells and the mechanism would allow escape
from autoimmunity. Receptor editing is more likely to result
from the rearranging mechanism that is not tightly regulated
and can go on or be reactivated without a specific need.
Properties of CDR3 in Xenopus
Composition
As mentioned above, CDR3 contributes most to the conforma-
tion of the antigen-combining site, and DH elements are the
source of most of CDR3 diversity. In Xenopus, DH segments can
potentially be used in all six reading frames if no stop codon
occurs. DH elements can be fused, a flexibility not shared with
mammalian DHs (13, 46, 58, 60). At the center of the antigen-
binding site, CDR3 amino acids form solvent exposed loops
rich in glycine and tyrosine. Accordingly, in mammals, IgH
CDR3s encoding glycines and tyrosines are selected for during
B-cell differentiation, whereas highly hydrophobic IgH CDR3s
are not selected (61). Similarly, Xenopus IgH CDR3 sequences
from different developmental stages are rich in hydrophilic and
loop-promoting residues (Fig. 5). IgH CDR3s encoding the gly-
cine-rich DH reading frame 1 of the two most frequently used
DH elements, DH1 and 10, are selected for during B-cell differen-
tiation. Interestingly, not a single strongly hydrophobic CDR3
sequence was found in Xenopus, even in the pre-B-cell popula-
tions of 5 and 7 day old larvae. In contrast to Xenopus, the
human, rabbit, mouse and chicken DH reading frame can pro-
duce highly hydrophobic sequences, but this rarely occurs in
peripheral B cells (62, 63). This indicates species-specific selec-
tion during evolution on the genomic sequences of Xenopus DH
elements. This might be linked to the pressure to rapidly
develop a functional immune system with a small number of
cells. The negative post-rearrangement selection of B cells due
to highly hydrophobic IgH CDR3 sequences, as observed in
mammals, is minimized in Xenopus, thus avoiding wastage, a fac-
tor always thought to play a role in shaping the immune system
of animals with small numbers of cells (64). The selection of a
germ-line DH sequence with a “compatible” sequence indepen-
dent of the reading frame could eliminate the need for early
selection steps such as those effected by the pre-B-cell receptor
in mammals (65). This might be relevant at early larval stages
of development, when the small number of B cells limits the
diversity of the B-cell repertoire.
Length
Most of the larval and young post-metamorphic Ig gene sam-
ples contained a CDR3 of 3–10 codons, two codons shorter
than adult CDR3s which were 5–12 codons long (66); the lat-
ter were shorter on the average than mammalian CDR3S, which
can reach up to 20 residues (58). Perhaps the shorter CDR3
generates a more limited spectrum of antigen-combining sites
(66). The reason for the short CDR3 in tadpole is in part due to
the lack of N diversity (13), which in adults is most likely to be
generated by terminal deoxynucleotidyl transferase (67).
Homology-based junction
In contrast to other DH and JH gene segments, DH1 and JH3 con-
tain overlapping sequences of two or three nucleotides at their
3' and 5' ends, respectively. In mammals it has been argued that
such short overlaps can direct the rearrangement, resulting in
redundant, so-called homology-based, V, D and J junctions (68,
69). Among 12 DH1–VH3 junctions from B-cell genomic DNA of
208 Immunological Reviews 175/2000
Du Pasquier et al · B-cell development in Xenopus
9–21 day old larvae, only two junctions, which also corre-
sponded to homology-based junctions, were identical. How-
ever, when cDNA of 23 day old larvae was analyzed, 12 out of
16 DH1–JH3 junctions were identical, and the dominant DH1–JH3
junctional sequence corresponded to one of the possible
homology-based junctions. The bias towards homology-based
junctions in cDNA sequences could therefore result from selec-
tion following rearrangement and leading to the presence of a
tyrosine at this position (44).
B cells at work
Antibody production in larvae and adults
Antibody responses can be elicted in Xenopus tadpoles (70, 71),
similar to tadpoles of other anurans such as the midwife toad
Alytes obstetricans (45), the bullfrog Rana catesbeiana (71, 72) and
Rana pipiens (73). From the moment the spleen becomes visible,
antibodies, mostly of the IgM type, can be detected (43). The
isotypes and the molecular nature of adult and larval Ig are
identical. IgM is first detected in the peritoneal fluid of larvae
at stage 49 (12 days) and IgG homolog at 15 days. IgX was seen
in liver plasma cells in tadpole but was not present in the gut,
which at this stage contains very few lymphoid cells and no B
cells. The ability of larvae to produce antibodies increased dur-
ing development from stage 50 to stage 59. The concomitant
injection of primed irradiated adult cells in addition to antigen
increased the amount of antibody made, especially the IgG
type. The anti-2,4-dinitrophenol (anti-DNP) antibodies pro-
duced by larvae were of lower affinity than those produced by
adults. Sibling larvae given additional primed adult cells pro-
duce more antibody but of the larval type (43, 74). Tadpole
and adult Xenopus, manipulated to be of comparable size by
blocking development with sodium perchlorate, exhibited
stage-specific antibody expression. The production of adult-
type higher affinity anti-DNP antibodies is independent of the
age and size of the individual and is concomitant with the com-
pletion of metamorphosis. The appearance of new antibody
specificities at such a time suggests that their expression occurs
with the cell turnover and renewal during a period of morpho-
logical change (47, 75). Specific memory, despite all the
changes, can cross metamorphosis (72). There is a relatively
higher membrane Ig expression in tadpoles compared with
adults. This could mean that plasma cell differentiation is not
optimal at early stages of development (76). Immunized and
A B
C D
Fig. 5. Xenopus IgH CDR3 sequences from different developmental stages. Amino acid sequence deduced from cDNA sequences of 5–7 day old larvae (A), DNA sequences of 15, 21, and 48 day old larvae (B), and cDNA sequences of immunized adults (C, D) (13, 78). The codons between the invariant Cys92 and Trp103 are shown. Residues encoded by nucleotides potentially derived from a DH element are shown in bold.
Immunological Reviews 175/2000 209
Du Pasquier et al · B-cell development in Xenopus
non-immunized individuals show different profiles of V region
expression. VHIII is expressed largely in non-experimentally
immunized animals. VHI is used preferentially in the anti-DNP
response as seen from protein sequences (77) and from the
cDNA sequences of immunized animals (76, 78).
Selection of somatic mutants during larval and adult immune
responses
Somatic mutations occur in Xenopus larvae or adults at a rate not
significantly lower than that observed in mammals (76, 78).
Why the antibody response is so poor compared with mam-
mals with little affinity maturation remains a mystery. Some
lines of evidence indicate that there is minimal selection of B
cells with higher affinity receptors. One line comes from com-
paring the actual ratio of replacement to silent mutations (R/S)
with the ratio expected from random mutation. R/S in CDR is
somewhat higher but not statistically significant. The other line
of evidence comes from the strong ratio of GC to AT base pairs
altered by mutations both in tadpoles and adults. It is possible
that selection of the mutants is limiting. To assess whether the
effect of selection can be observed in a system not involving Ig
genes we have used a tabulation of the spectra of spontaneous
mutations in seven different systems involving the loss of the
enzymes hypoxanthine guanine phosphoribosyltransferase and
adenine phosphoribosyltransferase in mammals (79). For each
system the log of the GC/AT mutation ratio was plotted against
the percentage of point mutations. In a loss of activity mutation
analysis the fraction of point mutations is a measure of the
stringency of selection. If almost total loss of activity is required
for mutants to get through the screen then the system being
analyzed will be enriched in other types of mutations. The log
of GC/AC mutation ratio correlates positively with the percent-
age of point mutations and hence correlates negatively with the
stringency of selection (79). Similarly, the mammalian lym-
phoid cell line 18.81, which mutates spontaneously without
antigen selection in vitro, also shows a profound GC bias (80,
81). In addition, the pattern of mutation seen in Xenopus is sim-
ilar to that found in mice deficient in the MSH2 mismatch
repair protein (82), as if the hypermutation pathway frequently
mutated G and C or as if the Xenopus MSH2-dependent pathway
that normally correlates G and C mismatches is not operational.
This is compatible with two non-mutually exclusive notions:
1) that hypermutation at the Ig locus is caused by a reduction
(qualitative or quantitative) of the error correction machinery
normally present in all cells rather than by the presence of addi-
tional error-prone repair system in B cells, and 2) that there is
minimal selection of high affinity B cells by antigen in Xenopus
due to the lack of germinal centers (see above) (59, 76, 78).
Switch
Some selection for higher affinity antibody occurs as immune
responses develop and progress in Xenopus. Under the influence
of T cells, heavy chain class switch occurs in Xenopus. cDNA
libraries made from immunized animals have higher ratios of
IgY/IgM clones than those from non-experimentally immu-
nized animals (13). The mechanism appears similar to that in
mammals, with loop excision and disappearance of the m locus
on the switched allele (83). The Xenopus Ig locus contains switch
regions that participate in the somatic transition from the
expression of one isotype to the other. When performed within
the first week of age, thymectomy in Xenopus prevents the switch
from IgM to IgY (84) but not from IgM to IgX (9). Low levels
of switching also occur in tadpoles, with IgX-positive cells hav-
ing been detected in the liver of day 30 tadpoles. IgY in tadpole
can be produced upon repeated immunization or after injec-
tion of adult syngeneic T cells (43). In addition, the switch to
IgY is highly temperature dependent in Xenopus. IgM switching
in animals at 19°C does not occur until 1 month after immuni-
zation. At 22°C, switch is easily detected after 21 days, and IgY
can be analyzed on isoelectric focusing gels. It is even more
readily induced at 27°C (57).
The switch region of IgM, S�, of about 5 kb has been iden-
tified in the JH–C� intron. S� contains 23 repeats each with a
length of approximately 150 bp. Each repeat consists of internal
smaller repeats and palindromic sequences, such as AGCT,
which they share with mammalian switch regions. A deletion
of the �-gene and joining of the S regions of � and occurs in
Xenopus B cells expressing IgX. Although not sequenced in its
totality, the switch region of IgX, S�� shows no sequence homol-
ogy with S� in its 3' portion and is characterized by 16 and 121
bp repeats and a high frequency of CATG, AGCA and TGCA pal-
indromes. Both IgM and IgXS regions are AT rich and not GC rich
like mammalian S regions. The majority of the recombinations
occur at positions (microsites) where a single strand DNA fold-
ing program predicts the transition from a stem to a loop struc-
ture. This feature is conserved in most mammalian switch junc-
tions, which points to the general existence and involvement of
microsites as a component that determines the recombination
breakpoint. The recombinogenic nature of the switch regions
is therefore linked to its structure rather than to its base com-
position, the repetitive occurrence of palindromes being essen-
tial for creating many microsites (83).
Conclusions
In Xenopus, all the major events involved in mammalian B-cell
differentiation occur (Fig. 6), consistent with homogeneity of
210 Immunological Reviews 175/2000
Du Pasquier et al · B-cell development in Xenopus
the immune system in vertebrates. However, results from
studying B-cell differentiation at very early developmental
stages in Xenopus, when the number of cells is small and perhaps
limiting, reveal differences that can alter interpretations of
events at the molecular level during early ontogeny in mam-
mals.
Among the many questions that remain unanswered is:
why is there minimal selection for antibody-combining site
mutations? This review reveals many gaps in our knowledge
about the genesis of the lymphoid organs and the cellular traffic
responsible for selection. This topic could be the focus of future
studies in Xenopus involving projects on chemokines and inter-
leukins. Research should move away from the major genetic
loci that have been studied so far, such as MHC, Ig and TCR, even
though much remains to be done in this area (for instance TCR
a, g, and d have still not been isolated).
Xenopus is an interesting model because of its ontogeny, but
its pseudotetraploid nature and its relatively long generation
time turn it into a genetic nightmare. With the prospect of fur-
ther developing the genetically simpler X. tropicalis, which also
takes only 8–10 months to reach sexual maturity, things may
soon change. In addition, Xenopus proves a good model for
transgenesis. Before transgenesis, nuclear transplantation mod-
els were once thought to be exploitable for performing single
lymphocyte genetics but were hampered by the fact that tad-
poles derived from lymphocyte nuclei did not live long enough
to allow immunological experiments (85, 86). The introduc-
tion of transgenic animals (87), especially using the genetically
simple X. tropicalis (88), can increase our possibilities and
should allow such experimental projects to regain momentum
in the future. This will probably require a painful and labor-
intensive adaptation to the new species; new libraries and
reagents will need to be created. However, a model will then be
available where molecular genetics and physiological aspects of
the immune system can be studied in a single species. Finally,
B cells do not develop in isolation. T cells and accessory cells
Fig. 6. The ontogeny of the B-cell compartment in Xenopus. [Modified from (2)]
Immunological Reviews 175/2000 211
Du Pasquier et al · B-cell development in Xenopus
should also be considered simultaneously. With the cloning of
TCR genes and preparation of new monoclonal antibodies,
more tools are available, and the prospect of comprehensively
studying the developing Xenopus immune system becomes more
realistic and feasible. The addition of lymphoid cell lines to the
arsenal of tools for the immunologist is welcome and has
already made possible the isolation and sequencing of two lym-
phocyte cell surface proteins and the elucidation of some
aspects of Ig gene rearrangement (16, 24, 89, 90).
References1. Du Pasquier L, Schwager J, Flajnik MF.
The immune system of Xenopus. Annu Rev Immunol 1989;7:251–275.
2. Du Pasquier L, Wilson M, Robert J. The immune system of Xenopus with special reference to B-cell development and immunoglobulin genes. In: Tinsley RC, Kobel HR, eds. The biology of Xenopus. Oxford: Clarendon Press; 1996. p. 301–313.
3. Horton JD, Horton TL, Ritchie P. The immune system of Xenopus: T cell biology. In: Tinsley RC, Kobel HR, eds. The biology of Xenopus. Oxford: Clarendon Press; 1996. p. 279–299.
4. Schwager J, Grossberger D, Du Pasquier L. Organization and rearrangement of immunoglobulin M genes in the amphibian Xenopus. EMBO J 1988;7:2409–2415.
5. Hadji-Azimi I, Michea-Hamzehpour M. Xenopus laevis 19S immunoglobulin. Ultrastructure and J chain isolation. Immunology 1976;30:587–591.
6. Schwager J, Mikoryak CA, Steiner LA. Amino acid sequence of heavy chain from Xenopus laevis IgM deduced from cDNA sequence: implicatons for evolution of immunoglobulin domains. Proc Natl Acad Sci USA 1988;85:2245–2249.
7. Amemiya CT, Haire RN, Litman GW. Nucleotide sequence of a cDNA encoding a third distinct Xenopus immunoglobulin heavy chain isotype. Nucleic Acids Res 1989;17:5388.
8. Mussmann R, Wilson M, Marcuz A, Courtet M, Du Pasquier L. Membrane exon sequences of the three Xenopus Ig classes explain the evolutionary origin of mammalian isotypes.Eur J Immunol 1996;26:409–414.
9. Hsu E, Flajnik MF, Du Pasquier L. A third immunoglobulin class in amphibians. J Immunol 1985;135:1998–2004.
10. Mussmann R, Du Pasquier L, Hsu E. Is Xenopus IgX an analog of IgA? Eur J Immunol 1996;26:2823–2830.
11. Hsu E, Schwager J, Alt FW. Evolution of immunoglobulin genes: VH families in the amphibian Xenopus. Proc Natl Acad Sci USA 1989;86:8010–8014.
12. Haire RN, Amemiya CT, Suzuki D, Litman GW. Eleven distinct VH gene families and additional patterns of sequence variation suggest a high degree of immunoglobulin gene complexity in a lower vertebrate, Xenopus laevis. J Exp Med 1990;171:1721–1737.
13. Schwager J, Bürckert N, Courtet M, Du Pasquier L. The ontogeny of diversification at the immunoglobulin heavy chain locus in Xenopus. EMBO J 1991;10:2461–2470.
14. Schriml LM, et al. Tyramide signal amplification (TSA)-FISH applied to mapping PCR-labeled probes less than 1 kb in size. Biotechniques 1999;27:608–613.
15. Haire RN, Ohta Y, Litman RT, Amemiya CT, Litman GW. The genomic organization of immunoglobulin VH genes in Xenopus laevis shows evidence for interspersion of families.Nucleic Acids Res 1991;19:3061–3066.
16. Du Pasquier L, Courtet M, Robert J. A Xenopus tumor cell line with complete Ig gene rearrangements and T-cell characteristics. Mol Immunol 1994;32:583–593.
17. Yu CC, Larijani M, Miljanic IN, Wu GE. Differential usage of VH gene segments is mediated by cis elements. J Immunol 1998;161:3444–3454.
18. Hsu E, Lefkovits I, Flajnik M, Du Pasquier L. Light chain heterogeneity in the amphibian Xenopus. Mol Immunol 1991;28:985–994.
19. Schwager J, Burckert N, Schwager M, Wilson M. Evolution of immunoglobulin light chain genes: analysis of Xenopus IgL isotypes and their contribution to antibody diversity. EMBO J 1991;10:505–511.
20. Stewart SE, Du Pasquier L, Steiner LA. Diversity of expressed V and J regions of immunoglobulin light chains in Xenopus laevis. Eur J Immunol 1993;23:1980–1986.
21. Haire RN, et al. A third Ig light chain gene isotype in Xenopus laevis consists of six distinct VL families and is related to mammalian � genes. J Immunol 1996;157:1544–1550.
22. Hsu E, Du Pasquier L. Studies on Xenopus immunoglobulins using monoclonal antibodies. Mol Immunol 1984;21:257–270.
23. Bhat NM, Kantor AB, Bieber MM, Stall AM, Herzenberg LA, Teng NN. The ontogeny and functional characteristics of human B-1 (CD5+ B) cells. Int Immunol 1992;4:243–252.
24. Jurgens JB, Gartland LA, Du Pasquier L, Horton JD, Gobel TW, Cooper MD. Identification of a candidate CD5 homologue in the amphibian Xenopus laevis. J Immunol 1995;155:4218–4023.
25. Hadji-Azimi I, Coosemans V, Canicatti C. B-lymphocyte populations in Xenopus laevis. Dev Comp Immunol 1990;14:69–84.
26. Barritt LC, Turpen JB. Characterization of lineage restricted forms of a Xenopus CD45 homologue. Dev Comp Immunol 1995;19:525–536.
27. Du Pasquier L, Hsu E. Immunoglobulin expression in diploid and polyploid interspecies hybrid of Xenopus: evidence for allelic exclusion. Eur J Immunol 1983;13:585–590.
28. Kobel HR, Du Pasquier L. Genetics of polyploid Xenopus. Trends Genet 1986;12:310–315.
29. Raff MC, Owen JJ, Cooper MD, Lawton ARD, Megson M, Gathings WE. Differences in susceptibility of mature and immature mouse B lymphocytes to anti-immunoglobulin-induced immunoglobulin suppression in vitro. Possible implications for B-cell tolerance to self. J Exp Med 1975;142:1052–1064.
30. Flajnik MF, Hsu E, Kaufman JF, Du Pasquier L. Biochemistry, tissue distribution and ontogeny of surface molecules detected on Xenopus hemopoietic cells. In: Miyasaka M, Trnka Z, eds. Differentiation antigens on lymphohemopoietic tissues. New York: M. Dekker; 1988. p. 387–419.
31. Du Pasquier L, Flajnik MF. Expression of MHC class II antigens during Xenopus development. Dev Immunol 1990;1:85–95.
32. Manning MJ. The effect of early thymectomy on histogenesis of the lymphoid organs in Xenopus laevis. J Embryol Exp Morphol 1971;26:219–229.
212 Immunological Reviews 175/2000
Du Pasquier et al · B-cell development in Xenopus
33. Zapata A, Amemiya CT. Phylogeny of lower vertebrates and their immunological structures. Curr Top Microbiol Immunol 2000;248:67–107.
34. Hsu E, Julius MH, Du Pasquier L. Effector and regulator functions of splenic and thymic lymphocytes in the clawed toad Xenopus. Ann Immunol 1983;3:277–292.
35. Greenhalgh P, Olesen CE, Steiner LA. Characterization and expression of recombination activating genes (RAG-1 and RAG-2) in Xenopus laevis. J Immunol 1993;151:3100–3110.
36. Koni PA, Flavell RA. Lymph node germinal centers form in the absence of follicular dendritic cell networks. J Exp Med 1999;189:855–864.
37. Baldwin WMD, Cohen N. A giant cell with dendritic cell properties in spleens of the anuran amphibian Xenopus laevis. Dev Comp Immunol 1981;5:461–473.
38. Zlotnik A, Morales J, Hedrick JA. Recent advances in chemokines and chemokine receptors. Crit Rev Immunol 1999;19:1–47.
39. Haynes L, Cohen N. Further characterization of an interleukin-2-like cytokine produced by Xenopus laevis T lymphocytes.Dev Immunol 1993;3:231–238.
40. Watkins D, Parsons SV, Cohen N. The ontogeny of interleukin production and responsitivity in the frog Xenopus. Thymus 1988;11:113–122.
41. Rollins-Smith LA, Blair P. Contribution of ventral blood island mesoderm to hematopoiesis in postmetamorphic and metamorphosis-inhibited Xenopus laevis. Dev Biol 1990;142:178–183.
42. Chrétien I, Marcuz A, Du Pasquier L. A ploidy marker to track lymphocytes after cell transfer between genetically identical or inbred Xenopus. In: Lefkovits I, ed. Immunology methods manual. London: Academic Press; 1997. p. 2381–2394.
43. Hsu E, Du Pasquier L. Ontogeny of the immune system in Xenopus. I. Larval immune response. Differentiation 1984;28:109–115.
44. Mussmann R, Courtet M, Du Pasquier L. Development of the early B-cell population in Xenopus. Eur J Immunol 1998;28:2947–2959.
45. Du Pasquier L. Ontogeny of the immune response in animals having less than one million lymphocytes: the larvae of the toad Alytes obstetricans. Immunology 1970;19:353–362.
46. Mussmann R. B-cell development in the amphibian Xenopus [PhD Dissertation]. University of Basel, Switzerland; 1996.
47. Du Pasquier L, Weiss N. The thymus during the ontogeny of the toad Xenopus laevis: growth, membrane-bound immunoglobulins and mixed lymphocyte reaction. Eur J Immunol 1973;3:773–777.
48. Hadji-Azimi I, Schwager J, Thiebaud C. B-lymphocyte differentiation in Xenopus laevis larvae. Dev Biol 1982;90:253–258.
49. Sollbach A, Marshall A, Yu C, Pennycook J, Wu GE. Understanding patterns of immunoglobulin gene rearrangements.Semin Immunol 1994;6:197–206.
50. Willems van Dijk K, Milner LA, Sasso EH, Milner EC. Chromosomal organization of the heavy chain variable region gene segments comprising the human fetal antibody repertoire. Proc Natl Acad Sci USA 1992;89:10430–10434.
51. Wu GE, Paige CJ. VH gene family utilization in colonies derived from B and pre-B-cells detected by the RNA colony blot assay. EMBO J 1986;5:3475–3481.
52. Kirkham PM, Schroeder HW Jr. Antibody structure and the evolution of immunoglobulin V gene segments. Semin Immunol 1994;6:347–360.
53. Pascual V, Verkruyse L, Casey ML, Capra JD. Analysis of Ig-H chain gene segment utilization in human fetal liver. Revisiting the “proximal utilization hypothesis”. J Immunol 1993;151:4164–4172.
54. Potter KN, Capra JD. Immunoglobulin variable region gene segments in human autoantibodies. In: Honjo T, Alt FW, eds. Immunoglobulin genes. London: Academic Press; 1995. p. 379–422.
55. Gu H, Kitamura D, Rajewsky K. B-cell development regulated by gene rearrangement: arrest of maturation by membrane-bound D� protein and selection of DH element reading frames. Cell 1991;65:47–54.
56. Wabl MR, Du Pasquier L. Antibody patterns in genetically identical frogs. Nature 1976;264:642–644.
57. Du Pasquier L, Wabl MR. Antibody diversity in amphibians: inheritance of isoelectric focusing antibody patterns in isogenic frogs. Eur J Immunol 1978;8:428–433.
58. Du Pasquier L, Schwager J. Immunoglobulin genes and B-cell development in amphibians. Adv Exp Med Biol 1991;292:1–9.
59. Du Pasquier L, Wilson M, Greenberg A, Flajnik MF. Somatic mutation in ectothermic vertebrates: musing on selection and origins. Curr Top Microbiol Immunol. 1998;229:199–216.
60. Du Pasquier L. Phylogeny of B-cell development. Curr Opin Immunol 1993;5:185–193.
61. Raaphorst FM, Timmers E, Kenter MJH, Van Tol MJD, Vossen JM, Schuurman RKB. Restricted utilization of germ-line VH3 genes and short diverse third complementarity-determining regions (CDR30) in human fetal B lymphocyte immunoglobulin heavy chain rearrangements. Eur J Immunol 1992;22:247–251.
62. Milili M, Schiff C, Fougereau M, Tonnelle C. The VDJ repertoire expressed in human pre-B-cells reflects the selection of bona fide heavy chains. Eur J Immunol 1996;26:63–69.
63. Nasman I, Lundkvist I. Evidence for oligoclonal diversification of the VH6-containing immunoglobulin repertoire during reconstitution after bone marrow transplantation. Blood 1996;87:2795–2804.
64. Du Pasquier L. Antibody diversity in lower vertebrates–why is it so restricted? Nature 1982;296:311–313.
65. Melchers F, Rolink A. B-lymphocyte development and biology. In: Paul WE, ed. Fundamental immunology. 4th ed. Philadelphia: Lippincott-Raven; 1999. p. 183–224.
66. Desravines S, Hsu E. Measuring CDR3 length variability in individuals during ontogeny. J Immunol Methods 1994;168:219–225.
67. Lee A, Hsu E. Isolation and characterization of the Xenopus terminal deoxynucleotidyl transferase. J Immunol 1994;152:4500–4507.
68. Feeney AJ. Predominance of VH-D-JH junctions occurring at sites of short sequence homology results in limited junctional diversity in neonatal antibodies. J Immunol 1992;149:222–229.
69. Gu H, Förster I, Rajewsky K. Sequence homologies, N sequence insertion and JH gene utilization in VHDJH joining: implications for the joining mechanism and the ontogenic timing of Ly1 B-cell and B-CLL progenitor generation. EMBO J 1990;9:2133–2140.
70. Kidder GM, Ruben LN, Stevens JM. Cytodynamics and ontogeny of the immune response of Xenopus laevis against sheep erythrocytes. J Embryol Exp Morphol 1973;29:73–85.
71. Du Pasquier L, Haimovich J. The antibody response during amphibian ontogeny. Immunogenetics 1976;3:381–391.
72. Haimovich J, Du Pasquier L. Specificity of antibodies in amphibian larvae possessing a small number of lymphocytes.Proc Natl Acad Sci USA 1973;70:1898–1902.
73. Marchalonis JJ. Ontogenetic emergence of immunoglobulins in Rana pipiens larvae. Dev Biol 1971;25:479–491.
Immunological Reviews 175/2000 213
Du Pasquier et al · B-cell development in Xenopus
74. Hsu E, Du Pasquier L. Ontogeny of the immune system in Xenopus II. Antibody repertoire differences between larvae and adults. Differentiation 1984;28:116–122.
75. Hsu E, Du Pasquier L. Changes in the amphibian antibody repertoire are correlated with metamorphosis and not with age or size. Dev Immunol 1992;2:1–6.
76. Wilson M, Marcuz A, Du Pasquier L. Somatic mutations during an immune response in Xenopus tadpoles. Dev Immunol 1995;4:227–234.
77. Brandt DC, Griessen M, Du Pasquier L, Jaton JC. Antibody diversity in amphibians: evidence for the inheritance of idiotypic specificities in isogenic Xenopus. Eur J Immunol 1980;10:731–736.
78. Wilson M, Hsu E, Marcuz A, Courtet M, Du Pasquier L, Steinberg C. What limits affinity maturation of antibodies in Xenopus-the rate of somatic mutation or the ability to select mutants? EMBO J 1992;11:4337–4347.
79. Zhang LH, Vrieling H, Zeeland AA, Jenssen D. Spectrum of spontaneously occuring mutations in the hprt gene of V79 Chinese hamster cells. J Mol Biol 1992;223:627–635.
80. Bachl J, Wabl M. An immunoglobulin mutator that targets GC. base pairs. Proc Natl Acad Sci USA 1996;93:851–855.
81. Meyer J, Jäck HM, Ellis N, Wabl M. High rate of somatic joint mutation in vitro and near the variable region segment of an immunoglobulin heavy chain gene. Proc Natl Acad Sci USA 1986;83:6950–6953.
82. Phung QH, et al. Increased hypermutation at G and C nucleotides in immunoglobulin variable genes from mice deficient in the MSH2 mismatch repair protein. J Exp Med 1998;187:1745–1751.
83. Mussmann R, Courtet M, Schwager J, Du Pasquier L. Microsites for immunoglobulin switch recombination breakpoints from Xenopus to mammals. Eur J Immunol 1997;27:2610–2619.
84. Turner RJ, Manning MJ. Thymic dependence of amphibian antibody responses. Eur J Immunol 1974;4:343–346.
85. Wabl MR, Brun RB, Du Pasquier L. Lymphocytes of the toad Xenopus laevis have the gene set for promoting tadpole development. Science 1975;190:1310–1312.
86. Du Pasquier L, Wabl MR. Transplantation of nuclei from lymphocytes of adult frogs into enucleated eggs: special focus on technical parameters. Differentiation 1977;8:9–19.
87. Kroll KL, Amaya E. Transgenic Xenopus embryos from sperm nuclear transplantations reveal FGF signaling requirements during gastrulation. Development 1996;122:3173–3183.
88. Amaya E, Offield MF, Grainger RM. Frog genetics: Xenopus tropicalis jumps into the future. Trends Genet 1998;14:253–255.
89. Du Pasquier L, Robert J. In vitro growth of thymic tumor cell lines from Xenopus. Dev Immunol 1992;2:295–307.
90. Robert J, Guiet C, Du Pasquier L. Lymphoid tumors of Xenopus laevis with different capacities for growth in larvae and adults.Dev Immunol 1994;3:297–307.